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AMERICAN
MALACOLOGICAL
BULLETIN

Fee 1988

——---
7 he!
oS Ae
ne @ yp Ry
oy af Fe
<0 bs)

VOLUME 6 NUMBER 1
CONTENTS | \ an
te £ ‘bs bout Cape, f
wag Pa Sy Sd

A comparison of growth rate between shallow water and deep water populations

of scallops, Placopecten magellanicus (Gmelin, 1791), in the Gulf of Maine.
DANIEL F. SCHICK, SANDRA E. SHUMWAY and MARGARET A. HUNTER

Genetic polymorphism in gastropods: a comparison of methods and habitat
scales. KENNETH M. BROWN and TERRY D. RICHARDSON

The mussels (Mollusca: Bivalvia: Unionidae) of Tennessee. LYNN B. STARNES
BGR URE et BOGAND cle cmMen Bie Meat a ee EME SPR he ooo oe Reman

Morphology of glochidia of Lampsilis higginsi (Bivalvia: Unionidae) compared
with three related species. D. L. WALLER, L. E. HOLLAND-BARTELS,

and L. G. MITCHELL
Research Note: A technique for trapping sandflat octopuses. JANET R. VOIGHT
49

Research Note: The need for quantitative sampling to characterize size
demography and density of freshwater mussel communities.

' ANDREW C. MILLER and BARRY S. PAYNE
SYMPOSIUM ON THE BIOLOGY OF THE POLYPLACOPHORA
Ancestors and descendents: relationships of the Aplacophora and
Rolypacophofa: AMELIENH: SCHELTEMAN! 8.5.24 Lae a Be ek ee PAN Le Pe 57
The gills of chitons (Polyplacophora) and their significance in molluscan
79

phylogeny. W. D. RUSSELL-HUNTER

A review of Caribbean Acanthochitonidae (Mollusca: Polyplacophora) with
descriptions of six new species of Acanthochitona Gray, 1821.
115

WILLIAM G. LYONS
Chitons (Mollusca: Polyplacophora) from the coasts of Oman and the
Arabian Gulf. PIET KAAS and RICHARD A. VAN BELLE
Sense organs in the girdle of Chiton olivaceus (Mollusca: Polyplacophora).
FRANZ PETER FISCHER, BRIGITTE EISENSAMER, CHRISTINA
Mize, INGHIO SINGER. Cam a.ye kee ee etits ikon. sae hee Ag bln waleksn ec oye hla saa 131
Sensory organs in the hairy girdles of some mopaliid chitons. ESTHER M. LEISE ............... 141
The ultrastructure of the aesthetes in Lepidopleurus cajetanus
Se ah META. Maer eek pe es. 2a 153
161
163

(Polyplacophora: Lepidopleurina). FRANZ PETER FISCHER

Financial Report
Announcements

i bias , *) .
ba te a ‘i
Pi So Pia ea
AMERICAN MALACOLOGICAL BULLETIN Geka
BOARD OF EDITORS POM aR Sh
EDITOR-IN-CHIEF MANAGING EDITOR | Ai
ROBERT S. PREZANT RONALD B. TOLL —
Department of Biology Department of Biology — ae
Indiana University of Pennsylvania University of the South cnt
Indiana, Pennsylvania 15705 Sewanee, Tennessee 37375 — ih
ASSOCIATE EDITORS IN . Bia A A
} F j yg iy OD i 7h GaN
MELBOURNE R. CARRIKER ROBERT ROBERTSON —_— ra A

College of Marine Studies

University of Delaware

Lewes, Delaware 19958

GEORGE M. DAVIS

Department of Malacology

The Academy of Natural Sciences
Philadelphia, Pennsylvania 19103

R. TUCKER ABBOTT
American Malacologists, Inc.
Melbourne, Florida, U.S.A.

JOHN A. ALLEN
Marine Biological Station
Millport, United Kingdom

JOHN M. ARNOLD
University of Hawaii
Honolulu, Hawaii, U.S.A.

JOSEPH C. BRITTON
Texas Christian University
Fort Worth, Texas, U.S.A.

JOHN B. BURCH
University of Michigan
Ann Arbor, Michigan, U.S.A.

EDWIN W. CAKE, JR.
Gulf Coast Research Laboratory
Ocean Springs, Mississippi, U.S.A.

PETER CALOW
University of Sheffield
Sheffield, United Kingdom

Department of Malacology —
The Academy of Natural Sciences

%

le a rennet 19103

RICHARD E. PETIT
Ex Officio
P.O. Box 30

North Myrtle Beach, South Carolina 29582

BOARD OF REVIEWERS |

JOSEPH G. CARTER
University of North Carolina
Chapel Hill, North Carolina, U.S.A.

ARTHUR H. CLARKE
Ecosearch, Inc.
Portland, Texas, U.S.A.

CLEMENT L. COUNTS, III
Coastal Ecology Research
University of Maryland

Princess Anne, Maryland, U.S.A.

THOMAS DIETZ
Louisiana State University

Baton Rouge, Louisiana, U.S.A. —

_ WILLIAM K. EMERSON

American Museum of Natural History
New York, New York, U.S.A.

DOROTHEA FRANZEN
Illinois Wesleyan University _
Bloomington, Illinois, U.S.A. _

VERA FRETTER

University of Reading
Berkshire, United Kingdom

ISSN 0740-2783

~ ROGER HANLON

W. D. RUSSELL-HUNTER
Department of Biology
Syracuse University ee
Syracuse, New York 13210 _

He

University of Texas i
Saeki Texas, USAR i simi 04°) 2

JOSEPH HELLER eae 9 ba
Hebrew oon of deligaaen
Jerusalem, Israel o

ROBERT E. HILLMAN
Battelle, New England sf
Duxbury, Massachusetts, US. A.

K. ELAINE HOAGLAND
Academy of Natural Sciences
Philadelphia, Pennsylvania, U.S.A.

. RICHARD S. -HOUBRICK ee

U.S. National Museum —

_ Washington, D.C., U.S.A.

VICTOR S. KENNEDY
University of Maryland
Cambridge, Maryland, U. S.A.

ALAN Gd: KOHN.) oa “e ie
University of Washington — a
Seattle, Wasting USA. pants

LOUISE RUSSERT KRAEMER
University of Arkansas
Fayetteville, Arkansas, U.S.A.

JOHN N. KRAEUTER
Baltimore Gas and Electric
Baltimore, Maryland, U.S.A.

ALAN M. KUZIRIAN
NINCDS-NIH at the
Marine Biological Laboratory

Woods Hole, Massachusetts, U.S.A.

RICHARD A. LUTZ
Rutgers University

Piscataway, New Jersey, U.S.A.

EMILE A. MALEK
Tulane University
New Orleans, Louisiana, U.S.A.

MICHAEL MAZURKIEWICZ
University of Southern Maine
Portland, Maine, U.S.A.

JAMES H. McLEAN
Los Angeles County Museum
Los Angeles, California, U.S.A.

ROBERT F. MCMAHON
University of Texas
Arlington, Texas, U.S.A.

ROBERT W. MENZEL
Florida State University
Tallahassee, Florida, U.S.A.

ANDREW C. MILLER
Waterways Experiment Station
Vicksburg, Mississippi, U.S.A.

BRIAN MORTON
University of Hong Kong
Hong Kong

JAMES J. MURRAY, JR.
University of Virginia
Charlottesville, Virginia, U.S.A.

RICHARD NEVES

Virginia Polytechnic Institute
and State University

Blacksburg, Virginia, U.S.A.

WINSTON F. PONDER
Australian Museum
Sydney, Australia

CLYDE F. E. ROPER
U.S. National Museum
Washington, D.C., U.S.A.

NORMAN W. RUNHAM

University College of North Wales

Bangor, United Kingdom

AMELIE SCHELTEMA
Woods Hole Oceanographic Institution
Woods Hole, Massachusetts, U.S.A.

ALAN SOLEM
Field Museum of Natural History
Chicago, Illinois, U.S.A.

DAVID H. STANSBERY
Ohio State University
Columbus, Ohio, U.S.A.

FRED G. THOMPSON
University of Florida
Gainesville, Florida, U.S.A.

THOMAS E. THOMPSON
University of Bristol
Bristol, United Kingdom

NORMITSU WATABE
University of South Carolina
Columbia, South Carolina, U.S.A.

KARL M. WILBUR
Duke University
Durham, North Carolina, U.S.A.

Cover. A dorsal view of Chiton fosteri Bullock from Oman. This and other chitons from Oman and the Arabian Gulf are discussed in a paper
by Kaas and Van Belle in this issue. This paper is one in a series of papers that appear herein as part of the proceedings of the 1987 American
Malacological Union Symposium on the Biology of the Polyplacophora.

THE AMERICAN MALACOLOGICAL BULLETIN is the official journal publication of the American Malacological Union.

AMER. MALAC. BULL. 6(1)

January 1988

A COMPARISON OF GROWTH RATE BETWEEN
SHALLOW WATER AND DEEP WATER POPULATIONS
OF SCALLOPS, PLACOPECTEN MAGELLANICUS (GMELIN, 1791),
IN THE GULF OF MAINE

DANIEL F. SCHICK, SANDRA E. SHUMWAY
AND
MARGARET A. HUNTER
DEPARTMENT OF MARINE RESOURCES
WEST BOOTHBAY HARBOR, MAINE 04575, U. S. A.

ABSTRACT

The rate of growth over several years has been compared between two shallow water (13-20 m)
and two deep water (170 m) populations of the sea scallop Placopecten magellanicus (Gmelin). Scallops
from one shallow water population were tagged and released in 1977 for fishermen to recapture. Addi-
tional scallops from a nearby population were tagged in 1978 for periodic retrieval and measurement
by divers over a Subsequent four year period. The deep water scallops were sampled periodically over
eight years (1976-1983), and their growth was measured through analysis of height-frequency of an
anomalously numerous year class spawned in 1975. The rate of growth of the offshore, deep water
scallops was found to be less than that of the inshore, shallow water scallops. The calculated max-
imum sizes attained, as determined by Ford-Walford plots are 150 mm for the shallow water popula-

tions and 110 mm for the deep water populations.

The giant scallop, Placopecten magellanicus (Gmelin),
is of considerable economic importance in eastern Canada
and the northeastern United States and has thus been the
subject of a number of growth studies (Stevenson, 1936;
Chaisson, 1949; Welch, 1950; Stevenson and Dickie, 1954;
Haynes, 1966; Merrill et a/, 1966; Naidu, 1969, 1975;
Jamieson, 1979; Posgay, 1979; Posgay and Merrill, 1979;
Ehinger, 1982; Serchuk et a/., 1982; Serchuk and Rak, 1983;
Choinard, 1984; Krantz et a/., 1984; Mohn et al., 1984; Mac-
Donald and Thompson, 1985; Roddick and Mohn, 1985). In
all but five of these studies, age was estimated and growth
was determined by the method of Merrill et a/. (1966) i.e. count-
ing shell rings and resilifer lines. Estimates of growth have
also been made by measuring increasing weight of somatic
tissue using either the adductor muscle (Haynes, 1966; Ser-
chuk and Rak, 1983; Mohn et a/., 1984) or whole somatic
tissue dry weight (MacDonald and Thompson, 1985). Krantz
et al. (1984) estimated growth by measuring temperature-
induced changes in 180/160 ratios in the scallop shell calcite.
Only Posgay (1963) and Naidu (1975) have measured growth
through tagging and recovery thus validating age determina-
tion in the species. None of the above studies include data
that show the effects of handling by repeated measurements

of shell growth of scallops in situ.

Variations in a number of allometric relationships with
depth for sea scallops have been observed (Schick et al.,
1987) and depth as a factor in scallop growth has been ad-
dressed previously by a number of authors (Caddy et a/., 1970;
Posgay, 1979; MacDonald and Thompson, 1985). Posgay
(1979) has noted a decrease in growth with increasing water
depth for scallops on Georges Bank at four ranges between
55 and 109 m. Caddy et a/. (1970), however, found little varia-
tion at five depth ranges between 55 and 144 m in the Bay
of Fundy. MacDonald and Thompson (1985) studied growth
at 10, 20, and 31 m off Newfoundland, Canada and found
decreasing growth with increasing depth.

In the present study, scallops were collected from two
shallow water populations (13-20 m) along the Maine coast
and two deep water populations (170 m) in the Gulf of Maine
to determine the extent to which the rate of growth varied with
depth.

MATERIALS AND METHODS

Specimens of the sea scallop, Placopecten
magellanicus, were collected from two shallow water and

American Malacological Bulletin, Vol. 6(1) (1988):1-8

1

2 AMER. MALAC. BULL. 6(1) (1988)

two deep-water locations in the Gulf of Maine for age and
growth determinations (Fig. 1). The shallow-water animals
were collected from Jericho Bay, Maine (44911.5’N, 68°30’S),
using an eight foot wide (2.4 m), three gang commercial drag
in tows of 10 minutes duration. Intact animals were measured
for shell height and length (Fig. 2) and then tagged by drill-
ing a hole through the upper valve over the byssal notch area
and inserting a Floy polyethylene spaghetti tag secured with
knots. Other workers have demonstrated that this method of
tagging is not harmful to the scallops (Posgay, 1963, 1981;
Naidu and Cahill, 1985). Scallops were held out of the water
for a maximum of three minutes during the tagging process
and were held in running seawater tanks prior to release to
minimize stress. In June 1977, 1000 scallops were broadcast
over commercial fishing grounds in Jericho Bay, an area with
depths ranging from 13-20 m identified by fishermen as be-
ing good scallop grounds. Tags and shells were returned by
fishermen over the next three years.

Another shallow water collection of scallops was ob-
tained in June 1978 by divers near Ringtown Island, Maine
(44°07.4'N , 68°29.4’S) an island just south of Jericho Bay.
These scallops were tagged, measured and placed by divers
in an area protected from dragging by rough bottom confor-
mation. They were subsequently recovered, remeasured and
released in November 1978, June 1979, December 1981 and
finally recaptured in June 1982 yielding the first scallop growth
data showing the effects of repeated handling.

In both shallow water scallop groups, growth was deter-
mined by measuring the increase in tangential shell height
of the left (top) valve between the time of tagging and time

+ ~—— +

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UNITED STATES CANADA

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45°|

MAINE

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Fig. 1. Location of shallow water and deep water sea scallop sampl-
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BSC

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Fig. 2. Sea scallop shell conformation and dimensions.

of recapture as well as measuring the height of rings formed
during that time (Naidu, 1975; Posgay, 1981). Shell height at
age was determined using the technique of Merrill et a/. (1966)
and a table of mean height-at-age was constructed for both
groups. Since ring formation occurs during the winter and
scallops spawn in the late summer, the age at first ring for-
mation is taken as 6 months with subsequent rings found at
18 months, 30 months, etc.

Deep water scallops from 170 m depth in the Gulf of
Maine were collected annually in August using a fine mesh,
32 ft. (9.8 m) chain footrope, semi-balloon otter trawl. Two deep
water locations provided continuous records between 1976
and 1983 except in 1980 when samples were unavailable.
These were: ~32 km (20 miles) South of Boothbay Harbor
(43°26.5'N, 69°33.3’S) and Jeffrey’s Basin (43904.25'N, 70°
11.33’S). Height frequency distributions over time were deter-
mined from shell heights measured to the nearest millimeter
(Figs. 3, 4). The increment in shell height of a predominant
year class (1975) from one year to the next was used as a
measure of growth. The shell heights are for August, and since
scallops spawn in August, the first height frequency is taken
as scallops at one year of age, with subsequent annual col-
lections representing scallops at 2 years, 3 years, etc. This
assumption of age is based on examination of the shells from
the first sample which showed one ring on each shell in-
dicating that they had survived one winter.

Controversy still exists over the most accurate method
of determining the correct age of scallops (Merrill et a/., 1966;
Krantz et a/., 1984). Unfortunately, our data do little to clarify

SCHICK ET AL.: GROWTH OF PLACOPECTEN 3

the situation. In four of the five shells of scallops retrieved
alive during the second year after the Jericho Bay tag
releases, there was one more ring than there should have
been between the file mark and the leading edge. These
shells should have contained one ring with growth before and
after the ring. Instead they contained two rings with growth
after the second ring. This happened in a small number of
returns, but did occur in four of the five specimens, indicating
perhaps more than a chance occurrence. No evidence of shell
margin chipping or other damage was apparent in the shells
to indicate the possibility of one of the rings being a shock
mark. None of the 93 first year tag returns showed any ring
formation. What caused the two rings to form in four of the
five living second year tag returns is unknown. Approximate-
ly 25 scallops from the general area that were tagged and
sequentially retrieved by divers over four years showed ex-
act correlation between numbers of years and number of
rings.

The offshore, deep water scallops, identified as essen-
tially one single year class, were sampled annually and a
height frequency histogram over time in the form of a
3-dimensional diagram was prepared. This histogram shows
a single size mode through time. Scallops from a 1980 col-
lection were read for shell ring structure by ten investigators.
Considerable variation in numbers of rings per shell occurred
between readers. The combined observed age distribution
of 43 scallops from the same year class read by 10 readers
was 5 at four years, 73 at five years, 237 at six years, 102 at
seven years, and 13 at eight years. The problems of ring iden-
tification and the variety of aids including corroboration by
resilifer marks have been addressed by Merrill et a/. (1966)
and still exist today. Also, the time from spatfall to the first
readable ring, usually around 25 mm, is open to question,
further clouding the relationship between a scallop’s size and
age.

Two problems arise in comparing the ring-structure
method of age determination with the 180 to 160 ratio in the
shell calcite method of Krantz et a/. (1984). First, the ratio work
was only performed on two shells. Second, age determina-
tion of these shells was by the method of Merrill et a/. (1966)
in which two readers agreed on age. There is enough dis-
crepancy in age determination between the two methods to
very probably negate any chance of the misreading of rings
causing the difference in age. Still, the unknowns of exact
age on both sides of the comparison leave the whole ques-
tion open to more definitive research being needed. Our first
year tag returns and our diver-retrieved aging results seem
to support the one ring one year theory, yet the second year
returns from the fishermen support the more than one ring
per year theory of Krantz et a/. (1984). Our offshore shells
seem to indicate one ring-one year, but the reader error or
perhaps the true ring number variation indicates the age
determination process is still inexact.

Ford-Walford plots were constructed from shell incre-
ment measurements taken from shallow water (Ringtown)
scallops to obtain an estimate of the von Bertalanffy growth
equation parameters Hoo and k. Ford-Walford plots were also
constructed for the deep water population at 32 km south

\20 =
<5 Sas
eStats eo y
5
gor
Ye
© 33

22 (%)

Fig. 3. Three-dimensional plot of shell height frequencies vs. time
from a deep water sea scallop population located 32 km south of
Boothbay Harbor, Maine.

Si

Fig. 4. Three-dimensional plot of shell height frequencies vs. time
from a deep water sea scallop population located west of Jeffrey’s
Ledge in the Gulf of Maine.

of Boothbay Harbor using the annual growth increments in
the predominant (1975) year class. The parameters Ho, k,
and t, for the von Bertalanffy growth equation, Ht} = Hoo
(1 - ek(to)), were determined from the height and age data
for both the shallow-water and deep-water populations. A com-
puter model was employed that uses an iterative process to
scan a grid of options for the parameters H-infinity, k and t,
for a least-squares fit given age-length data (Allen, 1966) and
calculates asymptotic confidence intervals for each parameter
(Ralston and Jennrich, 1978). All calculations were carried out
on an IBM 370 computer.

4 AMER. MALAC. BULL. 6(1) (1988)

Measuring growth by ring deposition in survivors of a
year class can be subject to a bias known as Lee’s
Phenomenon (Lee, 1912) where the results depend on selec-
tion factors in mortality of that year class. Selection factors
can favor slower growing scallops by killing off the faster grow-
ing individuals at a higher rate than the slower growing,
smaller individuals. If this occurs, then the mean size at age
of the survivors will be smaller than the mean size at age of
the year class without the selecting factor and the resulting
lag in mean size at age will bias the growth curve. Fishing
mortality is such a factor. If the fishing gear selection is for
taking larger individuals, the smaller scallops of any one year
class will be more likely to survive, producing a growth curve
that tails off faster than it should. In terms of von Bertalanffy
parameters, the iterative process could be forced to select
an H-infinity that is lower than it should be and that will force
the selection of a k value that is higher than it should be.
Due to the possibility of Lee’s Phenomenon from a variety
of factors, fishing mortality, handling of scallops, etc., it is
dangerous to look at only one parameter from a von Bertalanf-
fy curve and state that it is higher or lower than the same
parameter from another curve and attach any significance to
the comparison since all three of the parameters are related
and interactive.

RESULTS

Growth rates for four populations of Placopecten
magellanicus were determined. Mean shell height at age was
calculated for each shallow-water population from the shell
ring measurements (Table 1) and the same was calculated
for each deep-water population from the height frequency of
the one predominant year class measured annually (Table 2).
The same data were used to determine the von Bertalanffy
growth parameters and the asymptotic confidence intervals
for all four populations (Table 3).

Ford-Walford plots for the Ringtown Island population
and for the 32 km south of Boothbay Harbor population in-
dicate an H-infinity of 150 mm and 110 mm respectively (Fig.
5) which agrees closely with the empirical data (Tables 1, 2).
Further evidence for the growth rate difference is illustrated

160

e)
‘o)

D
{e)

Shallow Deep

Water Water
1498.8 \06.6
0301 0.328

H+! SHELL HEIGHT (mm)
@
[e)

£
oO

0 20 40 60 80 100 120 40 160
H, SHELL HEIGHT (mm)

Fig. 5. Shallow water and deep water Gulf of Maine sea scallop
growth: Ford-Walford plots and derived von Bertalanffy parameters.

in figure 6 which shows the von Bertalanffy growth curves for
each of the four populations. Note that the Ringtown Island
population gives a slightly lower curve than does the Jericho
Bay population, but both are higher than the curves for the
two deep-water populations.

In the von Bertalanffy growth equation, the parameters
H-infinity and k are inversely related. At the 32 km south of
Boothbay Harbor station, a heavy fishery for scallops in that
area cropped off the larger scallops starting in 1981, making
what appeared to be the predominant year class in the last
two years’ length-frequencies artificially low. An attempt to
separate the year classes in the length-frequencies for
1982-1983 by NORMSEP (Hasselblad, 1966) failed, so the
predominant bump was used in toto for both years. When the
iterative process in Allen’s (1966) fit of von Bertalanffy
parameters to the data was attempted, it found a better fit with

Table 1. Age-at-height key for shallow water scallops. Data generated from measured rings. Heights are given in mm.

LOCATION 0.5 15 25 35
Jericho Bay 11.4 39.3 63.7 88.7
Ringtown Island 71.0 90.7

AGE IN YEARS

45 5.5 65 75 85 9.5
110.2 123.5 135.7 — — —
101.1 110.0 122.3 128.2 136.5 140.0

Table 2. Age-at-height key for deep-water scallops. Data generated from height-frequency data. Heights are given in mm.

AGE IN YEARS
LOCATION 1.0 20 3.0 40 5.0 6.0 70 80
32 km South 27.2 51.4 64.1 759 — —_— 101.0 103.6

Boothbay Harbor
W. Jeffreys Ledge 259 41.2 575

_ 90.6 96.7 103.1 _

SCHICK ET AL.: GROWTH OF PLACOPECTEN S)

Table 3. Least squares regressions of Von Bertalanffy parameters for scallops from 4 locations in the Gulf
of Maine. Values are + 1 Asymptotic Confidence Interval.

Location Depth (m) K To Years fitted
Shallow Water

Jericho Bay 25 248 + 479 0.13 + 0.036 0.17 + 095 1-8

Ringtown Island 15 148 77 0.27 + 0.059 0.10 + 0.494 1-9
Deep Water

S. Boothbay Harbor 170 16 + 37 0.28 + 0025 -001 + 0.103 1-8

W. Jeffrey’s Ledge 174 223 + 353 0.09 + 0.019 -0.37 + 0.110 1-7

the lower H-infinity, and this raised the k value. The von Ber-
talanffy parameters (Table 3) and the curve (Fig. 6) for the
20 miles south of Boothbay Harbor scallop data reflects this
with a more rapid rise (higher k) in early years and with a tail-
ing off (lower H-infinity) in later years compared to the Jef-
frey’s Basin data.

The Ringtown Island scallops were measured by re-
peatedly collecting them, bringing them to the surface,
measuring them and returning them to the bottom. This
amount of handling could have retarded their growth in the
years during the measurements. Chapman (pers. comm.) and
Naidu (pers. comm.) have both indicated that handling, even
slight handling in an aquarium situation, can retard shell
deposition. The von Bertalanffy curve for the Ringtown Island
scallop data shows good growth in the early years, before the
scallops were caught, and much slower growth in the last few
years. The salient point is that even with the possibility of
retarded growth due to repeated handling, the Ringtown Island
scallop growth is stiil greater than the growth of the deep-water
scallops. Note that the shallow-water growth curves are based
on annual increments beginning at six months of age whereas
the deep water growth curves are based on annual increments
beginning at one year of age. The scallops sampled ranged
in age from one to nine years.

DISCUSSION

The data presented here clearly demonstrate a marked

€
(=
— 160 . Yericho Bay (inshore)
= ee
i} uae
@ 140 eee _, Ringtown (inshore )
o zon i
a 120 a __-© W. Jeffreys Ledge (offshore)
® 100 = pec § BBH (offshore)
wn at ane
80 aa ieee Ot
60 a a
ee
40
20
ie) eit ni n 1 1 n 4 n i EY a en
fo) ' 2 3 4 5) € 7 8 9 10
AGE (Years)

Fig. 6. Von Bertalanffy growth curves for two shallow water and two
deep water sea scallop populations in the Gulf of Maine.

difference in growth rate between shallow water and deep
water scallop populations and also represents the first in situ
study of the effects of handling on growth of scallops. Our
data are in general agreement with previously published
growth data for Placopecten magellanicus and further sup-
port the theory that growth is depth dependent, with increas-
ing depth representing deteriorating environmental suitability.

A number of authors have reported on growth rates in
Placopecten magellanicus and their results are briefly sum-
marized here only as they apply to the present study. Welch
(1950), in one of the earliest studies of growth in this species,
reported height-at-age measurements for scallops from
Jericho Bay, the same area used in the present investigation.
His reported average values of 46.3 mm for the second ring
and 142.2 mm for the ninth ring for Jericho Bay scallops are
indistinguishable from the data presented here 35 years later.
Naidu (1969, 1975) monitored growth in a northern, shallow
water population of P magellanicus and reported H-infinity
values for three locations ranging from 140 to 161 and k values
for the same areas ranging from 0.19 to 0.27. These are in
close agreement with those reported here for shallow water
(Table 3). Posgay and Merrill (1979) reported height-at-age data
for scallop samples collected by the National Marine Fisheries
Service from Georges Bank and the mid-Atlantic region dur-
ing the period 1958-1965. Their values for scallops collected
along the Maine coast ranged from 40 mm at the second ring
to approximately 143 mm at the ninth ring. Again, this com-
pares favorably with our reported values of 39.3 and 140 mm
for the second and ninth rings respectively. In a more recent
study, Serchuk et a/. (1982) presented data for scallops from
the Gulf of Maine, Georges Bank and the mid-Atlantic bight.
These authors showed that the scallops collected from a
range of depths in the Gulf of Maine had a smaller mean size-
at-age than those from Georges Bank or the mid-Atlantic bight
during the first seven years.

MacDonald (1984) and MacDonald and Thompson
(1985), in more recent studies of Placopecten magellanicus,
summarized the existing information on growth in this species
and compared growth at three depths in two areas off New-
foundland, Canada as well as off St. Andrews, New Bruns-
wick, Canada and off New Jersey, U. S. A. with collaborative
data from various depths in three other locations off New-
foundland. They showed that growth varied with depth at all
but one location, that growth was variable between locations,
and that growth differences could be attributed to measured

6 AMER. MALAC. BULL. 6(1) (1988)

differences in food and temperature.

Naidu (1975) found a latitudinal shift in the rate of
growth such that the environment in the more northerly loca-
tions produced larger maximum-size scallops with a slower
growth rate than their more southerly counterparts. Mac-
Donald and Thompson (1985), however, found no such lati-
tudinal differences in growth rate, nor did Serchuk et al. (1982).
While MacDonald and Thompson (1985) demonstrated slower
growth rates in deep water locations than in shallow water
areas at their two sampling locations in Newfoundland, they
found no differences in the Bay of Fundy near St. Andrews.
The differences in growth rates recorded between sampling
areas were attributed to variations in environmental

parameters. It seems most likely that environmental variables
such as temperature, depth and most importantly, food avail-
ability, account for the observed differences in scallop growth
rates. Choinard (1984) reported the lowest growth rates to date
for Placopecten magellanicus and attributed these slow rates
to the extreme water temperature regime of the Northumber-
land Strait. Jamieson (1979) also reported low growth rates
although not as low as Choinard for a different region of the
Northumberland Straight, Central Strait, and also attributed
his findings to the wide range of water temperatures in the
area. Von Bertalanffy parameters for the sea scallop reported
in the literature are summarized by location, author and date
in Table 4.

Table 4. Parameters of the von Bertalanffy growth equation (Hi = Ho (1-e “k(tto)) for the sea scallop, Placopecten magellanicus.

Hoo k to r2 Location Source
Newfoundland, Canada
Port-au-Port Bay Naidu, 1975
152 0.21 -0.48 Boswarlos
161 0.19 -0.88 West Bay
140 0.27 0.11 Fox Is. River
Sunnyside MacDonald and Thompson, 1985
176.5 0.19 0.55 0.97 10 m
165.5 0.20 0.63 0.97 20 m
158.4 0.16 0.10 0.97 31m
Dildo MacDonald and Thompson, 1985
174.5 0.19 0.66 0.97 10 m
168.2 0.19 0.37 0.96 20 m
1478 0.22 0.74 0.97 31m
Terre Nova N.P. MacDonald and Thompson, 1985
163.1 0.24 1.26 0.90 10 m
151.1 0.22 0.37 0.94 20m
146.0 0.17 —0.88 0.92 31m
Colinet MacDonald and Thompson, 1985
158.6 0.18 0.54 0.96 6m
160.1 0.19 0.72 0.96 16m
Northumberland St., PE.I., Canada
Tormantine Bed Choinard, 1984
103.76 0.37 0.6734 July
108.83 0.326 0.4636 November
114.8 0.276 —0.276 Central Strait Jamieson, 1979
Bay of Fundy, N.B., Canada
St. Andrews MacDonald and Thompson, 1985
166.9 0.21 0.51 0.96 10m
166.0 0.21 0.53 0.98 31m
170.2 0.19 0.20 0.97 76 m
174.3 0.22 -—1.238 Gulf of Maine, U.S.A. Serchuk et al., 1982
Georges Bank, U.S.A.
148.9 0.26 1.0 Georges Bank Posgay, 1962
145.5 0.38 1.5 Georges Bank Brown et al., 1972
146.4 0.35 1.4 Georges Bank Posgay, 1976
143.6 0.37 1.0 Georges Bank Posgay, 1979
152.5 0.34 -1.454 Georges Bank Serchuk et al., 1982
161.38 0.178 1.195 Georges Bank Roddick and Mohn, 1985
146.5 0.30 1.32 Northeast Peak Posgay, 1959
141.8 0.28 1.0 Northern Edge Posgay, 1959
1518 0.30 -1.126 Mid-Atlantic Bight, U.S.A. Serchuk et al., 1982

SCHICK ET AL.: GROWTH OF PLACOPECTEN He

The effects of environmental variables on growth rate
in scallops have most recently been demonstrated by Mac-
Donald and Thompson (1985). They showed, quite convinc-
ingly, that the growth rates were directly related to a combina-
tion of temperature and food availability with low temperature
and low food levels producing the smallest and slowest grow-
ing scallops. Posgay (1979) showed a decrease in mean size
at age with depth at Eastern Georges Bank. Over four depth
ranges from 55 m to 100 m, Posgay showed a decrease in
mean size at the fifth ring from 119 mm to 94 mm. Caddy et
al. (1970) however, did not show significant differences with
depth over the five depth ranges from 55 to 144 m in the Bay
of Fundy. The mean size of scallops at the fifth ring in their
study showed no significant differences between samples.
Since these two studies represent different sample sites, it
is likely that the differences in growth rates can again be at-
tributed to environmental differences. It is interesting to note
here that the Bay of Fundy scallops in both studies, Caddy
et al. (1970) and MacDonald and Thompson (1985), were the
animals that showed no differential growth with depth pro-
bably due to hydrographic homogeneity created by strong tidal
mixing.

Studies are currently underway to assess the available
food rations for the two populations being studied here. It
would appear that the reported slow growth rates and smaller
size-at-age for the deep-water scallops are primarily due to
a lack of suitable food items (Shumway et a/., 1987).

ACKNOWLEDGMENTS

The authors are indebted to S. K. Naidu, B. A. MacDonald and
R. L. Stephensen for reading an earlier version of the manuscript
and for offering valuable criticisms. We are also indebted to K.
Pinkham for collection of offshore scallop samples and to the Depart-
ment of Marine Resources research dive team for the collection of
inshore samples. The authors wish to thank C. Crosby for his technical
expertise and organizational determination and M. Dunton for his quiet
ability to get things done.

LITERATURE CITED

Allen, K. R. 1966. A method of fitting growth curves of the von Ber-
talanffy type to observed data. Journal of the Fisheries Research
Board of Canada 23:163-179.

Brown, B. E., Parrack, M. and D. D. Flescher. 1972. Review of the
current status of the scallop fishery in International Commis-
sion for the Northwest Atlantic Fisheries (ICNAF) Division 5 Z.
ICNAF Research Document 72/113, Serial No. 2829, 13 pp.

Caddy, J. F., Chandler, R. A. and E. |. Lord. 1970. Bay of Fundy Scallop
Surveys 1966 and 1967 with observations on the commercia!
fishery. Technical Report No. 168, Fisheries Research Board
of Canada. 9 pp.

Chaisson, L. P. 1949. Report of scallop investigations and explora-
tions in the southern Gulf of St. Lawrence-1949. Fisheries
Research Board of Canada, Manuscript Report, Biological Sta-
tion No. 395.

Choinard, G. 1984. Growth of the sea scallop (Placopecten
magellanicus) on the Tormentine Bed, Northumberland Strait.
Canadian Atlantic Fisheries Scientific Advisory Committee
(CAFSAC) Research Document 84/61, 16 pp.

Ehinger, R. E. 1978. Seasonal energy balance of the sea scallop
Placopecten magellanicus from Narragansett Bay. Master’s
Thesis, University of Rhode Island, Kingston, Rhode Island.
88 pp.

Hasselblad, V. 1966. Estimation of Parameters for a mixture of Nor-
mal Distributions. Technometrics 8(3):431-444.

Haynes, E. B. 1966. Length-weight relation of the sea scallop
Placopecten magellanicus (Gmelin). International Commission
for the Northwest Atlantic Fisheries (ICNAF) Research Bulletin
3:1-17.

Jamieson, G. S. 1979. Status and assessment of Northumberland
Strait scallop stocks. Fish and Marine Service Technical Report
No. 904. 12 pp.

Krantz, D. E., Jones, D. S. and D. F. Williams. 1984. Growth rates of
the sea scallop, Placopecten magellanicus, determined from
the 180/160 record in shell calcite. Biological Bulletin 167:186-199.

Lee, R. M. 1912. An investigation into the methods of growth deter-
mination in fishes. Conseil Permanent International Pour L’ex-
ploration de la Mer, Publication de Circonstance 63:1-35 pp.

MacDonald, B. A. 1984. The partitioning of energy between growth
and reproduction in the giant scallop Placopecten magellanicus
(Gmelin). Doctoral Dissertation, Memorial University of New-
foundland. 202 pp.

MacDonald, B. A. and R. J. Thompson. 1985. Influence of temperature
and food availability on the ecological energetics of giant
scallop Placopecten magellanicus (Gmelin). |. Growth rates of
shell and somatic tissue. Marine Ecology Progress Series
25:279-294.

Merrill, A. S., Posgay, A. and F. E. Nichy. 1966. Annual marks on shell
and ligament of sea scallop (Placopecten magellanicus) Fishery
Bulletin 65(2):299-311.

Mohn, R. K., Robert, G. and D. L. Roddick. 1984. Georges Bank
scallop stock assessment - 1983. Canadian Atlantic Fisheries
Scientific Advisory Committee (CAFSAC) Research Document
84/12, 27 pp.

Naidu, K. S. 1969. Growth, reproduction and unicellular endosym-
biotic alga in the giant scallop Placopecten magellanicus
(Gmelin) in Port au Port Bay, Nfld. Master’s Thesis, Memorial
University of Newfoundland. 101 pp.

Naidu, K. S. 1975. Growth and population structure of a northern
shallow-water population of the giant scallop, Placopecten
magellanicus (Gmelin). International Council for the Explora-
tion of the Sea (ICES) Council Meeting 1975/K: 37, 17 pp.

Naidu, K. S. and F. M. Cahill. 1985. Mortality associated with tag-
ging in the sea scallop, Placopecten magellanicus (Gmelin).
Canadian Atlantic Fisheries Scientific Advisory Committee
(CAFSAC) Research Document 85/21, 10 pp.

Posgay, J. A. 1962. Maximum yield per recruit of sea scallops. In-
ternational Council for the Exploration of the Sea (ICES) Coun-
cil Meeting 1979/K:27:1-5.

Posgay, J. A. 1981. Movement of tagged sea scallops on Georges
Bank. Marine Fisheries Review 43(4):19-25.

Posgay, J. A. and A. S. Merrill. 1979. Age and growth data for the
Atlantic coast sea scallop, Placopecten magellanicus. National
Marine Fisheries Service-North East Fishery Center (NMFS-
NEFC) Laboratory Reference Document 79-58.

Ralston, M. L. and R. I. Jennrich. 1978. DUD, a derivative-free
algorithm for nonlinear least squares. Technometrics 1:7-14.

Roddick, D. L. and R. K. Mohn. 1985. Use of age-length informa-
tion in scallop assessments. Canadian Atlantic Fisheries
Scientific Advisory Committee (CAFSAC) Research Document
85/37, 16 pp.

Schick, D. F., S. E. Shumway and M. Hunter. 1987. Allometric rela-
tionships and growth in Placopecten magellanicus: the effects

8 AMER. MALAC. BULL. 6(1) (1988)

of season and depth. Malacological Review (in press).

Serchuk, F. M., Wood, P. W. and R. S. Rak. 1982. Review and
assessment of the Georges Bank, Mid-Atlantic and Gulf of
Maine Atlantic sea scallop (Placopecten magellanicus)
resources. National Marine Fishery Service (NMFS)/Woods
Hole Reference Document No. 82-06, 132 pp.

Serchuk, F. M. and R. S. Rak. 1983. Biological characteristics of
offshore Gulf of Maine sea scallop populations: Size distribu-
tions, shell height-meat weight relationships and relative fecun-
dity patterns. National Marine Fishery Service (NMFS)/Woods
Hole Reference Document No. 83-07, 42 pp.

Shumway, S. E., R. Selvin and D. F. Schick. 1987. Food resources
related to habitat in the scallop Placopecten magellanicus

(Gmelin, 1791): A qualitative study. Journal of Shellfish
Research (in press).

Stevenson, J. A. 1936. The Canadian scallop: Its fishery, life history,
and some environmental relationships: Master’s Thesis, Uni-
versity of Western Ontario, London, Ontario, Canada. 191 pp.

Stevenson, J. A. and L. M. Dickie. 1954. Annual growth rings and
rate of growth of the giant scallop Placopecten magellanicus
(Gmelin) in the Digby area of the Bay of Fundy. Journal of the
Fisheries Research Board Canada 11(5):660-671.

Welch, W. 1950. Growth and spawning characteristics of the scallop
in Maine waters. Master’s Thesis, University of Maine, Orono.
95 pp.

Date of manuscript acceptance: 30 March 1987

GENETIC POLYMORPHISM IN GASTROPODS: A COMPARISON

OF METHODS AND HABITAT SCALES

KENNETH M. BROWN
AND
TERRY D. RICHARDSON
DEPARTMENT OF ZOOLOGY AND PHYSIOLOGY
LOUISIANA STATE UNIVERSITY
BATON ROUGE, LOUISIANA 70803, U. S. A.

ABSTRACT

We compare genetic differentiation in gastropods at two habitat scales, using two methodologies.
For the pond pulmonate, Lymnaea elodes (Say), we present data on the degree of genetic variance
for life histories by comparing variation in traits among full sib groups reared in a common field en-
vironment, for two source populations (one vernal, one permanent pond). For the same two popula-
tions, as well as a third in another vernal pond, we also present data on allozyme polymorphism. Finally,
we contrast genic polymorphism occurring over a much broader habitat scale, using published literature
on allozyme polymorphism found respectively in terrestrial, freshwater, and marine environments, and
for snails having selfing, outcrossing, or parthenogenetic mating systems.

We found more genetic variation for life history traits in Lymnaea elodes occurring in a vernal
pond, as variation among sib groups was significant for 5 out of 6 traits measured, versus only 2 out
of 6 in a population from a permanent pond. We thus found that unpredictable habitats can favor greater
levels of genetic variation in life histories. In contrast, genic polymorphism was similar in all three ponds,
with from 27 to 33% of loci polymorphic, and mean heterozygosity ranging only from 8 to 10%. Genetic
similarity was high for the two vernal ponds and lower for the more distant permanent pond. Divergence
in heterozygosity did occur across broader habitat categories, with lower mean heterozygosity for snails
in terrestrial habitats, and self-fertilizers in particular possessing significantly lower heterozygosity within
this habitat. The literature survey also indicated more work on allozyme variation is needed in par-
ticular for freshwater pulmonates, and we suggest such work along with further work on variation in

polygenic characters like life histories.

Although many studies have looked at variation in
bioenergetics, life histories, and shell structure among popula-
tions of freshwater snails (See reviews in Russell-Hunter, 1978;
Russell-Hunter and Buckley, 1983; and McMahon, 1983), lit-
tle is known of the genetic basis of variation among or within
populations for these traits (Brown, 1983). In contrast, quite
a bit is known, from studies of allozyme variation, about levels
of genic polymorphism in freshwater as well as marine snails
(see reviews in Clarke et a/., 1978; Berger, 1983; Nevo et al.,
1983; Selander and Ochman, 1983). No studies have as
yet attempted to study both the genetic basis for variation in
life histories and genic polymorphism among and within the
same populations of a species. We present such data on varia-
tion among sib groups of snails for a number of life history
patterns, contrasting them among two populations of the pond
snail Lymnaea elodes (Say). One population is from a vernal,
the other a permanent pond in northern Indiana. For these

same two populations, as well as a second vernal pond, we
also present data on allozyme variation. To determine if trends
in mean heterozygosity appear at a broader habitat scale than
between populations, we also review the available literature
on allozyme variation in freshwater, marine, and terrestrial
gastropods. Within each of these broad habitat categories,
we further divide populations as to their breeding systems,
including selfing, outcrossing, and parthenogenetic reproduc-
tion to determine whether these reproductive modes have any
broad effect on polymorphism.

For Lymnaea elodes, several studies have concen-
trated on proximal factors affecting population dynamics, in-
cluding density dependence (Eisenberg, 1970), habitat pro-
ductivity and permanence (Hunter, 1975; Brown et al/., 1985),
and water temperature (Brown, 1979). Brown (1985) used
transfer experiments to show that most variation among
populations in life histories was due to habitat productivity.

American Malacological Bulletin, Vol. 6(1) (1988):9-17

9

10 AMER. MALAC. BULL. 6(1) (1988)

However, lack of divergence among populations could also
be due to pronounced phenotypic and/or genetic variation
within populations. For this reason we decided, using snails
from both populations, to rear offspring from different sib
groups in the same field environment to determine the scale
of differences in life histories occurring across sib groups.

We decided to study allozyme polymorphism in the
same populations of Lymnaea elodes for two reasons. First,
genic polymorphism is probably an independent estimator of
genetic variation in populations, when compared to genetic
variation in phenotypes such as life histories (Lewontin, 1984).
Second, little is known of allozyme variation among and within
populations of lymnaeid snails or, for that matter, most other
freshwater pulmonates. In contrast, allozyme variation has
been studied in detail within and among populations of
freshwater prosobranchs (Chambers, 1978, 1980; Selander
et al., 1978; Dillon and Davis, 1980; Karlin et a/., 1980;
Selander and Ochman, 1983; Dillon, 1984). Marine
gastropods, which are almost exclusively prosobranchs, have
also been studied extensively for allozyme polymorphism.
Studies have included many species of Cerithium (Ritte and
Pashton, 1982; Lavie and Nevo, 1986), Crepidula (Hoagland,
1985; Woodruff et a/., 1986), Littorina (Ward and Warwick,
1980), marsh snails such as Nassarius (Gooch et al., 1972),
thaidids (Garton, 1984; Garton and Stickle, 1985), and a deep-
water species, Bathybembix bairdii (Dall) (Siebenaller, 1978).

The genetic structure of terrestrial gastropods is
perhaps the best known, with studies ranging from slugs (Foltz
et al., 1982a, b, 1984) to a large number on shelled species
such as Cerion, Cepaea, and Partula (see references listed
in appendix). We have done the first survey to look explicitly
for differences in polymorphism among gastropods across
broad habitat categories, although several reviews have con-
sidered the effect of mating systems on heterozygosity
(Selander and Kaufman, 1975; Nevo et al., 1983; Selander
and Ochman, 1983).

METHODS

THE SPECIES AND HABITATS

Lymnaea elodes is a common algivore in temporary
ponds and marshes in the northern tier of states in the United
States as well as Canada (Brown, 1979). Its life cycle length,
fecundity, and shell growth are determined for the most part
by habitat productivity and adult snail density (Eisenberg,
1970; Hunter, 1975; Brown, 1985). Adults reproduce in late
spring and early summer, and juveniles and some adults
estivate over late summer (if the pond dries) and winter (Brown
et al., 1985). Lymnaea elodes can be eliminated from more
permanent habitats (e.g. lakes) by fish predators (Brown and
DeVries, 1985). Snails for this study were collected from a ver-
nal pond (Pond A, Brown, 1982) and a more permanent pond
(Pond F, Brown, 1982) for experiments assessing variation
among sib groups. The vernal pond usually dries by early July
and the more permanent pond has water at least until August,
after oviposition has been completed. Snails were also col-

lected from a second vernal pond (Pond B, Brown, 1982) for
the electrophoretic analyses. This pond usually dries in late
July. The three ponds also differ in food levels, with the per-
manent pond having the greatest periphyton productivity
(Brown et al., 1985). All ponds are located in Noble County,
Indiana within 30 km of Crooked Lake Biological Station,
33 km NW of Fort Wayne.

VARIATION AMONG SIB GROUPS

The permanent pond was selected as a common rear-
ing site for both populations since higher food levels would
not limit egg production (Brown et al., 1985) and snails would
be able to complete their life cycles before pond drying. Thir-
ty juveniles were collected from the temporary pond and
twenty-five juveniles from the permanent pond in early spring
1981. These snails were placed singly in flow-through con-
tainers in Pond F and reared through their entire life cycle.
They were paired with a snail from the same source pond for
a two week interval when they were between 12 mm and
14 mm shell length to allow outcrossing (lymnaeid snails, like
other pulmonates, are hermaphroditic). Companion snails
were removed before experimental snails reached 16 mm, the
smallest recorded shell length at maturity in these popula-
tions (Brown et al., 1985), so that egg laying would not be con-
founded between the two individuals. Since Lymnaea elodes
outcrosses preferentially (Brown, 1979), we have assumed
snails did not self-fertilize. At weekly intervals, we measured
snails and removed all egg cases. Eggs and hatched juveniles
were kept over winter in 6/ aquaria at 13°C in the laboratory
to retard growth and maturation. In spring 1982, we placed
an average of 3.5 full sib offspring per temporary pond parent,
and 3.0 full sib offspring for each permanent pond parent back
in the permanent pond. The same rearing methods were used
for offspring as for their parents in the preceding season. A
more detailed account of the rearing methods is given in
Brown et al. (1985).

Six life history traits were measured for each offspring:
(1) shell length at maturity; (2) relative age at maturity (days
since start of experiment); (3) clutch size (average number
of eggs per mass); (4) total fecundity; (5) shell length at death;
(6) relative age at death (days since start of the experiment).
Exact ages at maturity and death were impossible to deter-
mine, as offspring were not separated in the laboratory by date
laid. The impact of differing growth rates of offspring over
winter in the laboratory was minimized by using initial shell
length as a covariate in an analysis of covariance (ANCOVA)
that assessed differences among sib groups in each of the
life history patterns.

ALLOZYME VARIATION

Approximately 150 snails were collected from each
pond and were frozen at —60°C until analysis when the foot
was ground in a chilled cell mill with 0.5 ml of deionized water.
Supernatant was absorbed onto wicks of Whatman #3 filter
paper and inserted into horizontal starch gels and subjected
to electrophoresis for 3-5 hours at 35-55 mA and 200 volts.
Gels were stained for the following enzyme systems: (1) acid
phosphatase (ACP); (2) esterases (EST); (3) aspartate amino-

BROWN AND RICHARDSON

transferase (AAT); (4) glucose phosphate isomerase (GPI); (5)
hexanol dehydrogenase (HEX); (6) leucine aminopeptidase
(LAP); (7) malate dehydrogenase (MDH); (8) mannose-6-
phosphate isomerase (MPI); (9) 6-phosphogluconate dehy-
drogenase (PGD); (10) sorbitol dehydrogenase (SDH); (11)
superoxide dismutase (SOD). All loci were examined for all
populations. Formulas for gel and electrode buffers, as well
as stains for these enzyme systems were taken from Shaw
and Prasad (1970), Selander et a/. (1971), Chambers (1980),
and Dillon and Davis (1980). Enzymes with multiple loci were
numbered by mobility (1=fastest).

Allelic and genotypic frequences were calculated us-
ing the BIOSYS-1 FORTRAN program (Swofford and Selander,
1981) which also calculated the percentage of loci polymor-
phic (using the criterion that a second allele must have a fre-
quency = 5%). Mean observed and expected heterozygosities
(over all loci) for each population and genetic distance indices
between populations were also calculated. BIOSYS-1 uses
the methods of both Rogers and Nei to calculate genetic
distance.

In the literature survey, we either used reported heter-
ozygosities, or calculated heterozygosity over all loci (including
monomorphic loci as 0% heterozygous) if raw data on allelic
or genotypic frequencies were reported. Each population was
then catalogued by habitat and reported mating system. If we
could not determine from the paper whether the gastropod
was a Selfer, outcrosser, or parthenogen, it was placed in the
facultative selfing category along with species reported as
having a mixed breeding strategy. Data were arc-sine trans-
formed and subjected to ANOVA. The ideal design would be
factorial, allowing us to look at the interactive effects of habitats
and breeding systems. Due to empty cells, we were con-
strained, however, to perform two separate one way analyses.
First we performed a oneway ANOVA over habitat categories.
Second, we performed a oneway ANOVA over breeding
systems within each habitat category. Duncan’s a posteriori
multiple range tests were used to compare means (at the 0.05
significance level) if the F statistic was significant.

RESULTS

VARIATION AMONG SiB GROUPS

For sib groups from the permanent pond, the covariate
(initial shell length) had significant effects on four of the six
life history traits (Table 1). In contrast, only two life history traits,
shell length at maturity and clutch size, showed significant
variation among sib groups. Variation among sib groups in
life history traits was much more obvious in the temporary
pond, with significant effects occurring for five of the six traits
(Table 2). Initial shell length, however, still had significant ef-
fects on the same five traits. Thus, the ANCOVA suggests
greater levels of genetic variation for life histories in the ver-
nal pond population, when snails are reared in a common
field environment. However, the inital size of individuals when
introduced to containers also has substantial effects on life
history variation.

: GASTROPOD POLYMORPHISM il

Table 1. Analysis of Covariance, with initial shell length as the
covariate, of six life history traits among sib groups from a perma-
nent pond. Values are F statistics. One asterik indicates significance
at the 0.05 level, two at the 0.01 level.

SOURCES OF VARIATION
Among Within

Traits Treatments Covariate Sib Groups Sib Groups
Degrees

Freedom 24 1 23 60
Age at

Maturity 13 6.6* <1

Shell length

at Maturity 2.9** 28.3** 1.8*

Clutch Size 3.0** 6.8* 2.9**

Total

Fecundity 1.3 3.1 1.2

Age at

Death 1.7* 6.2* 1.5

Shell Length

at Death <1 <1 <1

Table 2. Analysis of Covariance, with initial shell length as the
covariate, for six life history traits among sib groups from a temporary
pond. Values are F statistics. One asterisk indicates significance at
the 0.05 level, two at the 0.01 level.

SOURCES OF VARIATION
Among Within

Traits Treatments Covariate Sib Groups Sib Groups
Degrees

Freedom 30 1 29 81
Age at

Maturity 6.2** 60.7** 43**

Shell length

at Maturity 3.6** 43.7** 2.3**

Clutch Size Silla 9.2** 2.9**

Total

Fecundity 2.8** 24.2** Zale

Age at

Death <1 <1 <1

Shell Length

at Death 2.4** 14.0** 2.0**

ALLOZYME POLYMORPHISM

Patterns in allozyme polymorphisms were consistent
across ponds. In all three populations, nine loci were
monomorphic: ACP; EST-1; EST-4; GPI; HEX; MPI; PGD; SDH;
SOD. In the snails from the first temporary pond (A), 26.7%
of the loci were polymorphic, including EST-3, AAT, LAP-1 and
MDH. In the second temporary pond, the LAP-2 locus was
also polymorphic, with one-third of the loci polymorphic overall
(Table 3). In the permanent pond, 26.7% of the loci were
polymorphic, namely EST-2, EST-3, LAP-1, and MDH (Table 3).

Mean observed heterozygosity was also similar in each

12 AMER. MALAC. BULL. 6(1) (1988)

Table 3. Allelic frequencies, mean expected and observed hetero-
zygosities, and proportion of total loci polymorphic for three pond
populations of Lymnaea elodes. N refers to number of individuals.

Second
Temporary Temporary Permanent

Locus’ Allele Pond Pond Pond
EST-2 A 0.99 1.0 0.32

B 0.01 0.0 0.68
EST-3 A 0.07 0.01 0.19

B 0.03 0.32 0.39

C 0.90 0.67 0.42
AAT A 0.85 0.63 1.0

B 0.15 0.37 0.0
LAP-1 A 0.55 0.01 0.02

B 0.32 0.35 0.29

Cc 0.13 0.64 0.69
LAP-2 A 0.99 0.95 0.99

B 0.01 0.05 0.01
MDH A 0.72 0.56 0.94

B 0.28 0.44 0.06
Mean Observed
Heterozygosity
(SE) 0.08(0.04) 0.10(0.04) 0.09(0.04)
Mean Expected
Heterozygosity
(SE) 0.10(0.05) 0.13(0.06) 0.11(0.06)
Percent of Loci
Polymorphic 26.7 33.3 26.7
N 150 150 150

Table 4. Pairwise estimates of genetic distance among populations
of the pond snail Lymnaea elodes. Nei’s “‘unbiased”’ indices are above
the main diagonal, Rogers’ indices below.

Second
Temporary Temporary Permanent

Pond Pond Pond
Temporary
Pond 0.0 0.03 0.07
Second
Temporary Pond 0.08 0.0 0.06
Permanent
Pond 0.14 0.12 0.0

of the populations and ranged from 8 to 10% (Table 3). Mean
expected heterozygosities calculated by BIOSYS-1 ranged
from 10 to 13%, indicating some degree of heterozygote defi-
ciency in all three ponds as is common in most molluscs. All
pair-wise comparisons of the three populations show levels
of genetic distance near 0.05 (Table 4), a level characteristic
of populations within the same species (Avise, 1976). Both
distance indices indicated snails from the permanent pond
to be more dissimilar from each of the 2 temporary ponds than
the 2 temporary ponds were from each other. This could be
due to lower levels of gene flow, since the permanent pond

is over 40 km from either of the temporary ponds, which are
separated by only 2 km.

Although percent of polymorphic loci and mean ob-
served heterozygosities were similar, there were some in-
teresting differences in allele frequencies among the 3 popula-
tions (Table 3). The 2 temporary ponds had similar allele fre-
quencies at EST-2 with allele A being most common. However,
in the permanent pond population allele B predominated with
a frequency of 0.68. For EST-3, allele C was most common
in the temporary pond A population, but dropped to 67% in the
second temporary pond population. In the permanent pond,
allele A was more common than in ponds A or B. AAT had
similar allelic frequencies in both temporary pond populations
with allele A declining from 0.85 to 0.63 in the second tem-
porary pond. The pond F population was monomorphic at this
locus. Ponds B and F were similar in allelic frequencies at
LAP-1 locus with allele C predominating. In contrast, in the
pond A population allele A was most common. LAP-2 did not
differ much in allelic frequencies among the 3 populations,
although with the = 5% criterion LAP-2 was polymorphic on-
ly in the pond B population. MDH differed somewhat in allelic
frequencies among the 3 populations although allele A was
always more common.

LITERATURE SURVEY

Genic polymorphism has been much more extensive-
ly studied in terrestrial gastropods, with over twice as many
populations represented than either of the other two habitat
categories (Table 5). Outcrossing appeared the most common
mating system in each habitat, and populations with obligate
selfing were found only in terrestrial snails. Parthenogenetic
populations have been studied only in freshwater snails (Table
5). The actual populations and heterozygosities used in the
analysis are given in the appendix.

Table 5. Overall mean heterozygosity for habitats and mating systems.
Means are weighted for sample size. Numbers in parentheses are
sample size.

Mating

System Terrestrial Freshwater Marine
Selfers 0.0 = =(6) — —
Outcrossers 0.089 (34) 0.106 (14) 0.173 (13)
Parthenogens — 0.207 (6) —
Facultative

Selfers 0.047 (10) 0.088 (1) 0.090 (2)
Overall mean 0.061 (50) 0.131 (21) 0.161 (15)

Mean observed heterozygosity was highly significant-
ly different among habitat types (F293 = 7.8; p <0.001). The
a posteriori test revealed that only terrestrial populations had
significantly lower average heterozygosity. Although average
heterozygosity can be lowest in terrestrial snails simply
because they alone possess selfing populations with no genic
polymorphism (Table 5), there appears also to be a general
trend, as terrestrial snails in both the outcrossing and

BROWN AND RICHARDSON: GASTROPOD POLYMORPHISM 13

facultative selfing categories had the lowest observed
heterozygosity of the three habitats. Within terrestrial
gastropods, there was, as might be expected, a highly signif-
icant difference in mean heterozygosity among mating
systems (F247 = 16.6, P<0.0001), and Duncan’s multiple
range test indicated selfers had a significantly lower average
heterozygosity. As also might be expected, outcrossing
gastropods had the highest average heterozygosity, and par-
tial selfers had intermediate heterozygosities. In both
freshwater and marine habitats there were no significant dif-
ferences among mating systems in average heterozygosity
(P>0.05). Finally, this study of polymorphism in Lymnaea
elodes reveals levels of heterozygosity just below the average
for outcrossing freshwater snails as a group (Table 5), in-
dicating the most probable mating system in these pond
populations is mixed.

DISCUSSION

The results of the full sib analyses indicate greater
levels of genetic variation for life history traits in snails drawn
from a vernal pond population. Perhaps the more unpredic-
table nature of this habitat has favored the maintenance of
genetic variation in life history traits. For example, the vernal
pond has extremely unpredictable drying dates from year to
year (Brown et a/., 1985). In wet years, juvenile recruitment
is good, and adult densities are high enough the next year
to depress fecundity by density dependence. In years with
little rainfall, the vernal pond dries so early that juvenile and
adult mortality are intense (Brown et al., 1985). If genetic varia-
tion in life histories provides a range of age at reproduction,
etc., then at least some individuals would successfully
reproduce regardless of the drying date, and genetic variance
for life history traits would be maintained. Interestingly, popula-
tions of pill clams in vernal ponds in Ohio also have more
genetic variation than populations in permanent ponds
(McCleod et a/., 1981; Burky, 1983). In addition, initial size of
individuals introduced to containers also affected life history
variation; as initial size increases so does clutch size, age
at maturity, shell length at maturity, and age at death (Brown
et al., 1985).

In contrast, the electrophoretic data indicate little dif-
ference in polymorphism between any of the populations of
Lymnaea elodes. Levels of polymorphism are very similar in
both of the vernal ponds, and essentially the same set of loci
vary in the permanent pond as well. Thus, interpretations on
levels of genetic variation within and among populations
based on the electrophoretic data do not agree with those
based on variation among full sibs in life history traits.
However, as Lewontin (1984) points out, when there is no
known functional relationship between the allozymes and
quantitative traits chosen (as in this case), there is no reason
to expect a pattern to emerge when comparing the two be-
tween populations. Even if a functional relationship exists be-
tween the allozymes and quantitative traits, the relative lack
of statistical power associated with gene frequency analyses
would require a prohibitively large sample size to detect

differences at the same level of statistical significance as the
quantitative traits. The allozyme polymorphism data do in-
dicate, however, little genetic differentiation among popula-
tions, similar to earlier transplant studies (Brown, 1985) sug-
gesting little genetic divergence among populations in life
histories. Compared to the average for all populations reported
in the literature, mean heterozygosity in these populations of
L. elodes is very near the value for outcrossing terrestrial
pulmonates, slightly less than the average for outcrossing
freshwater snails (again mostly dioecious prosobranchs) and
much less than dioecious prosobranch marine snails.
Therefore, these populations of L. elodes probably have a
mixed breeding system, with some inbreeding occurring, if
for no other reason than the fact that populations go through
bottlenecks when ponds dry early (Brown et a/., 1985). Also,
previous studies of mollusc populations indicate GPI, MPI,
PGD, SOD, and HEX are virtually always polymorphic (Clarke
et al., 1978; Selander and Ochman, 1983). Interestingly, these
loci were monomorphic in these 3 populations of L. elodes,
possibly due to the recurrent bottlenecks.

However, the literature survey indicated there were dif-
ferences in heterozygosity over broader habitat categories
than these pond populations of Lymnaea elodes. Terrestrial
pulmonates, regardless of the mating system, have the lowest
heterozygosities. This could be due to the nature of their
habitats. Terrestrial micro-environments hospitable to snails
(the proper temperature and humidity, etc.) might be expected
to be more patchily distributed than those in aquatic or marine
habitats (Russell-Hunter, 1983). Furthermore, terrestrial snails
are relatively immobile and might self-fertilize more than most
have considered. Effective population sizes could therefore
be low and inbreeding might occur, lowering levels of polymor-
phism (but for an exception see discussion in Cain, 1983).
One would expect that freshwater populations, due to their
seasonal nature, could again experience frequent bottlenecks,
resulting in lower levels of polymorphism than marine popula-
tions where many species also have widely dispersed,
planktonic larvae. Indeed, mean heterozygosity in freshwater
populations was intermediate to terrestrial and marine values.
In each of the habitats, outcrossers had the highest and
facultative selfers or selfers the lowest heterozygosity, as
would be expected. However, since many authors originally
classified populations as selfers only because of low
heterozygosity, these results could be somewhat circular.
Freshwater parthenogens, interestingly, had the highest
average heterozygosity. This suggests the existence of
apomictic clones within these populations, as is also seen
in parthenogenetically reproducing water fleas (Lynch, 1984)
or brine shrimp (Browne et al., 1984).

However, the interpretation of the literature survey was
confounded by a number of gaps in the data available on
genic polymorphism in gastropods. For example, is selfing
a common reproductive mode in other habitats besides ter-
restrial ones? Although the ability to self might be advan-
tageous in terrestrial habitats because of the patchy nature
of the proper microenvironments and consequent low den-
sities of conspecifics, we cannot be certain that predominately
selfing populations also occur in either freshwater or marine

14 AMER. MALAC. BULL. 6(1) (1988)

habitats, but have as of yet not been studied. Similarly, one
wonders if parthenogens occur in other habitats besides
freshwater. Vail (1978) reports that parthenogenesis is more
frequent in upriver viviparid populations, where densities are
low and chances of meeting males infrequent. If this is the
advantage for parthenogens in freshwater, why have not par-
thenogens evolved (or more likely been studied) in terrestrial
habitats since terrestrial snails are so patchily distributed?
Finally, the available data are confounded by taxonomic bias.
Most of the terrestrial snails are hermaphroditic pulmonates,
capable of self-fertilization, wnereas most of the aquatic and
marine populations are dioecious prosobranchs.

Overall, this study points to the need for more work
on levels of genetic variation in gastropods. We need further
studies on the degree of genetic variation in polygenic traits
like life histories both among and within populations. Contrasts
between temporary and permanent ponds, as well as other
important environmental parameters, would also be welcome.
Russell-Hunter (1983) suggests the need for incorporation of
factors like feeding niche (grazers vs. detritivores), reproduc-
tive modes (viviparity vs. oviviparity), possession of planktonic
larvae, and colonization ability as well. We also need to fill
in the gaps present in studies of allozyme variation, even
though existing work is more thorough than studies on varia-
tion in polygenic traits. In particular, more work is needed on
the degree of allozyme polymorphism among and within
populations of freshwater pulmonates. Although we know
much about variation in life histories and secondary produc-
tion among populations of freshwater pulmonates (See reviews
in Russell-Hunter, 1978; McMahon, 1983), much less is known
of the underlying genetic variation for these traits among and
within the same populations.

ACKNOWLEDGMENTS

We would like to gratefully acknowledge support from NSF
grant 81-03539 to the senior author, without which this work could
not have been done. Bonnie Leathers and Dennis DeVries helped
with the field work and electrophoresis. D. Pashley and D. Foltz read
and commented on an earlier version of the manuscript.

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Johnson, M. S., B. Clarke and J. Murray. 1977. Genetic variation and
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Karlin, A. A., V. A. Vail and W. H. Heard. 1980. Parthenogenesis and
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Lavie, B. and E. Nevo. 1986. Genetic diversity of marine gastropods:
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Lewontin, R. C. 1984. Detecting population differences in quantitative
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Livshits, G., L. Fishelson and G. S. Wise. 1984. Genetic similarity
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Lynch, M. 1984. The genetic structure of a cyclical parthenogen.
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McCleod, M. J., D. J. Hornbach, S. |. Guttman, C. M. Way and A.
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McCracken, G. F. and P. F. Brussard. 1980. The population biology
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Mulley, J. C. 1981. Polymorphism in the gastropod Austrocochlea con-
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Nevo, E., C. Bar-El and Z. Bar. 1981. Genetic structure and climatic
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Date of manuscript acceptance: 12 July 1987

16

AMER. MALAC. BULL. 6(1) (1988)

Appendix 1. Terrestrial gastropods. Observed heterozygosity and mating system (? or mixed = facultative selfing).

SPECIES

Milax sowerbyi (Férussac)

M. budapestensis (Hazay)
Limax maximus L.

L. pseudoflavus Evans

L. marginatus Muller
Deroceras caruanae (Pollonera)
D. reticulatum (Muller)

Milax gagates (Draparnaud)
Limax tenellus Muller
Deroceras agreste (L.)

Arion ater ater L.

a. rufus L.

. lusitanicus Mabille

. subfuscus (A) (Draparnaud)
. subfuscus (B)

. circumscriptus Johnston

. Silvaticus Lohmander

. hortensis Ferussac

. intermedius Normand

. distinctus Mabille

. owenii Férussac

Cerion bendalli Pilsbry and Vanatta
Deroceras laeve (Muller)

Helix aspera (Muller)

Rumina decollata (L.)
Sphincterochila aharonii (Kobelt)
S. cariosa (Oliver)

S. fimbriata (Bourguignat)

S. prophetarum (Bourguignat)
S. zonata (Bourguignat)
Theba pisana (Muller)

Partula gibba Bruguiere

P. mirabilis Crampton

P. olympia Crampton

P otaheitana Férussac

P suturalis Pfeiffer

P. taeniata Morch

Achatina fulica Bowdich
Bradybaena similaris Férussac
Cerion incanum (Burch and Kim)
Triodopsis albolabris (Say)
Xerocrassa seetzeni (Pfeiffer)
Cepaea nemoralis (L.)

C. hortensis (Muller)

C. sylvatica (Draparnaud)
Helix pomatia (L.)

Otala lactea Muller

O. vermiculata Muller
Oxychillas cellarius (Muller)
Nymphophilus minckleyi Taylor
Anguispira alternata (Say)

DDDEBDEREDRDEB

MATING
SYSTEM

outcross
outcross
outcross

?
outcross
outcross
outcross
outcross
outcross
selfer
mixed
outcross
outcross
outcross
mixed
selfer
selfer
outcross
selfer
outcross
outcross
outcross
mixed

?
selfer

?
outcross
outcross

?

?

ff
selfer
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross

STUDY

Foltz et al/., 1984

Foltz et al/., 1984

Foltz et al/., 1984

Foltz et al., 1984

Foltz et a/., 1984

Foltz et al., 1984

Foltz et al., 1984

Noble, unpubl.

Noble, unpubl.

Noble, unpubl.

Foltz et al., 1982a

Foltz et al., 1982a

Foltz et al., 1982a

Foltz et al., 1982a

Foltz et al., 1982a

Foltz et a/., 1982a

Foltz et a/l., 1982a

Foltz et al/., 1982a

Foltz et a/., 1982a

Foltz et al., 1982a

Foltz et al., 1982a

Woodruff, 1975

Foltz et al., 1982b

Selander and Kaufman, 1975
Selander and Kaufman, 1975
Nevo et al., 1983

Nevo et al., 1983

Nevo et al., 1983

Nevo et al., 1983

Nevo et al., 1983

Nevo et al/., 1981

Johnson et al., 1977
Johnson et al., 1977
Johnson et al., 1977
Johnson et al., 1977
Johnson et al., 1977
Johnson et al., 1977
Selander and Ochman, 1983
Selander and Ochman, 1983
Woodruff, 1978

McCracken and Brussard, 1980
Nevo, 1978

Jones et a/., 1980

Selander and Ochman, 1983
Selander and Ochman, 1983
Jarvinen et a/., 1976
Selander and Ochman, 1983
Selander and Ochman, 1983
Selander and Ochman, 1983
Selander and Ochman, 1983
Selander and Ochman, 1983

BROWN AND RICHARDSON: GASTROPOD POLYMORPHISM

Appendix 2. Freshwater gastropods. Observed heterozygosity and mating system.

SPECIES

Goniobasis vanhyningiana Goodrich

. floridensis (Reeve)

. dickinsoni Clench and Turner
. athearni Clench and Turner

. albanyensis Lea

. curvicostata (Reeve)
Melanoides tuberculata (Muller)
M. tuberculata (Muller)
Campeloma decisa (Say)

C. decisa (Say)

Lymnaea elodes

Biomphilaria straminea (Dunker)
B. glabrata (Say)

B. havanensis (Pfeiffer)

B. alexandria (Ehrenberg)
Campeloma geniculum (Conrad)
C. parthenum Vail
Potamopyrgus jenkinsi Smith
Viviparous contectoides (Binney)
Physa heterostropha (Say)
Helisoma trivolvis Say

QDHAADYD

Ho

0.031
0.077
0.066
0.182
0.184
0.078
0.306
0.111
0.095
0.033
0.088
0.082
0.30
0.091
0.068
0.250
0.375
0.138
0.112
0.171
0.136

MATING
SYSTEM

outcross
outcross
outcross
outcross
outcross
outcross
outcross
parth
parth
parth
mixed
outcross
outcross
outcross
outcross
parth
parth
parth
outcross
outcross
outcross

STUDY

Chambers, 1980

Chambers, 1980

Chambers, 1980

Chambers, 1980

Chambers, 1980

Chambers, 1980

Livshits et a/., 1984

Livshits et a/., 1984
Selander et a/., 1978
Selander et al., 1978

This study (all populations)
Woodruff et a/., 1985
Woodruff et a/., 1985
Woodruff et a/., 1985
Woodruff et a/., 1985

Karlin et a/., 1980

Karlin et a/., 1980

Selander and Ochman, 1983
Selander and Ochman, 1983
Selander and Ochman, 1983
Selander and Ochman, 1983

Appendix 3. Marine gastropods. Observed heterozygosity and mating system.

SPECIES

Adalaria proxima (Alder and Hancock)

Onchidoris muricata (Muller)
Thais haemastoma L.

T. lamellosa (Gmelin)
Cerithium scabridum Philippi
C. rupestre (Risso)
Crepidula onyx Sowerby

C. adunca Sowerby

C. fornicata (L.)
Austrocochlea constricta Fisher
Bathybembix bairdii (Dall)
Cerithium scabridum

C. caeruleum Sowerby
Nassarius obsoletus (Say)

Littorina rudis (Dautzerberg and Fisher)

L. arcana Ellis

Ho

0.082
0.059
0.106
0.017
0.168
0.035
0.161

0.052
0.045
0.168
0.162
0.620
0.635
0.166
0.153
0.132

MATING
SYSTEM
outcross
outcross
outcross
outcross

?

?
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross
outcross

STUDY

Havenhand et a/., 1986
Havenhand et a/., 1986
Garton, 1984

Garton and Stickle, 1985
Lavie and Nevo, 1986
Lavie and Nevo, 1986
Woodruff et a/., 1986
Woodruff et a/., 1986
Hoagland, 1985

Mulley, 1981

Siebenaller, 1978

Ritte and Pashtan, 1982
Ritte and Pashtan, 1982
Gooch et al., 1972

Ward and Warwick, 1980
Ward and Warwick, 1980

THE MUSSELS (MOLLUSCA: BIVALVIA: UNIONIDAE)
OF TENNESSEE

LYNN B. STARNES
UNITED STATES FISH AND WILDLIFE SERVICE
UNITED STATES DEPARTMENT OF THE INTERIOR
WASHINGTON, D. C. 20240, U. S. A.
AND
ARTHUR E. BOGAN
DEPARTMENT OF MALACOLOGY
THE ACADEMY OF NATURAL SCIENCES
PHILADELPHIA, PENNSYLVANIA 19103, U. S. A.

ABSTRACT

The unionid fauna that occurs within the political boundaries of the State of Tennessee is re-
viewed. The fauna reported from the Tennessee, Cumberland, Conasauga and Mississippi river
drainages is compared and discussed. There are 155 unionid taxa (species and subspecies) that cur-
rently occur or that have been reported historically from the state.

The State of Tennessee, because of the physiographic
diversity and discrete drainages encompassed by its boun-
daries, has one of the most diverse mussel faunas in North
America. The state’s molluscan fauna is enriched by virtue
of having four major river drainages: Mississippi, Tennessee,
Cumberland and Conasauga (Coosa River system) (Fig. 1).
Bickel (1968) listed 133 unionid taxa from Tennessee but in-
cluded only the fauna from the Tennessee and Cumberland
rivers. A total of 155 taxa have now been recorded from the
state. While the unionid fauna from the Tennessee and
Cumberland rivers has been historically documented and
periodically evaluated, the unionid fauna from the Mississip-
pi River and its direct tributaries in Tennessee, as well as the
Conasauga River, has only recently been described.

The vast majority of the unionid fauna is associated
with big river habitat. Pollution, channelization, commercial
harvest, impoundments and other modifications, have great-
ly reduced the extent of suitable riverine habitat, curtailing
distribution of many species. Of the 24 unionid species listed
by the U. S. Fish and Wildlife Service as threatened or en-
dangered, 18 (75%) occur in Tennessee (Hatcher and Ahl-
stedt, 1982; Bogan and Parmalee, 1983). Most of these
species are endemic to the Tennessee and Cumberland rivers
(Table 1).

This presentation reviews literature, archaeological and
unpublished museum records of the unionid fauna in the State
of Tennessee. An in depth analysis of each Tennessee unionid

species that involves taxonomy, shell description, distribution
and related data is currently under preparation by Dr. Paul
W. Parmalee, McClung Museum, University of Tennessee,
Knoxville.

RELEVANT FAUNAL STUDIES

Of the four major river drainages, the Tennessee River
unionid fauna is the most thoroughly studied. Pilsbry and
Rhoads (1897), Coker and Boepple (1912), Ortmann (1918),
Brown and Pardue (1980), Pardue (1981) and Dennis (1984)
described the unionid fauna in the upper Tennessee River
tributaries. Parmalee and Klippel (1984) documented the fauna
of the Tellico River, a tributary to the Little Tennessee River.
Bogan and Starnes (1983) discussed the Little River unionid
fauna. Hickman (1937) surveyed the Clinch River below Nor-
ris Dam, prior to the dam’s completion. Bates and Dennis
(1978) and Ahlstedt (1984) discussed the current status of the
unionid fauna of the Clinch River. Dennis (1981) summarized
some early historical and certain recent unionid data for the
Powell River. Ortmann (1925) described the fauna of the Ten-
nessee River and its tributaries in northern Alabama and
southern Tennessee. Isom (1972) reported the freshwater
bivalve fauna at the Nickajack Dam Site. Ortmann (1924)
described the fauna of the Duck River. Subsequently, van der
Schalie (1939, 1973), Isom and Yokley (1968) and Ahlstedt
(1981) documented drastic declines in the mussel fauna of
the Duck River. The Elk River was surveyed by Remington

American Malacological Bulletin, Vol. 6(1) (1988):19-37

19

20 AMER. MALAC. BULL. 6(1) (1988)

RIVER RIVER ELK

TENNESSEE RIVER

!
!
I

ALABAMA

T U Cc K Y 72
°
Ze NA
aga
2Ze° s
GS S, HOLSTON R
Y POMS ig i aaa
7
°
Ne
CANEY °
FORK v7
EMORY PRENCH N CAR
RIVER py. SROADR 7
& -
ca ee iy ie TEN
x / w NESse
Se Rive sste fj” re
Ow ° _—_s—
as -"
pee pp Ge ae
s
\ 4 S CAR
4 \,
\ CONASAUGA ~
i RIVER \
s
. GEORGIA
a
\ 0 125 250
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scale

Fig. 1. Map showing the major tributary rivers to the Cumberland, Tennessee and Mississippi rivers in the State of Tennessee.

and Clench (1925), Ortmann (1925), Isom et a/. (1973) and
Ahistedt (1983). Isom (1969) compared mussel faunas col-
lected in 1965 from the Tennessee River with those recorded
prior to impoundment. Scruggs (1960) and Isom and Gooch
(1986) made similar pre and post-impoundment comparisons.
Yokley (1972) compared the ecology and stocks of species
in Kentucky Reservoir. The Tennessee Valley Authority (TVA)
Cumberlandian Mollusk Conservation Program, detailed col-
lections from the Clinch, Powell, Nolichucky, Holston, Elk,
Duck and Buffalo rivers (Ahlstedt, 1986).

Unionids of the Cumberland River system in Tennessee
were studied by Wilson and Clark (1914), Neel and Allen
(1964), Isom et al. (1979), Parmalee et a/. (1980), Clarke (1981,
1985), Call and Parmalee (1982), Schmidt (1982), Sickel (1982),
Starnes and Bogan (1982) and Stansbery et a/. (1983). The
fauna in the Cumberland River appears similar to that of the
Tennessee River, but has not been as thoroughly surveyed
and future work could uncover significant differences.

Of the 87 mussel taxa recorded from the Tennessee
River, the 69 taxa recorded from the Duck River, and the 78
taxa recorded from the Cumberland River, Ortmann (1924)
considered 45 of these to be unique to the Tennessee and
Cumberland rivers and referred to them as ‘“‘Cumberlandian ’’.
Ortmann (1925) defined the downriver limits of the Cumber-
landian fauna to be Clarksville, Tennessee, on the Cumberland
River; Muscle Shoals, Alabama, on the Tennessee River; and
between Columbia and Centerville on the Duck River. Below
these limits, Interior Basin molluscan species replaced the
Cumberlandian species. Ortmann later liberalized these limits,
suggesting that some Cumberlandian species had emigrated
into the Ohio River as well as into the Interior Basin.

Reports of unionids from the Mississippi River
tributaries in Tennessee have been limited to Ortmann (1926a)
and van der Schalie and van der Schalie (1950). Recent col-
lections from the Hatchie River (D. Manning, pers. comm.)

suggest a diverse fauna. With the exception of the Hatchie
River, direct Mississippi River tributaries in Tennessee have
suffered extensive channelization resulting in major altera-
tions of their biological communities and a significant reduc-
tion of the unionid fauna.

The mussel fauna of the Conasauga River located in
the southeast corner of Tennessee is relatively unknown with
Hurd (1974), van der Schalie (1981) and museum records pro-
viding the only information on this northern Coosa River
tributary.

TAXONOMY

Table 1 lists unionid taxa found in the Tennessee and
Cumberland rivers in Tennessee. A comparison is made of
the nomenclature used by Bickel (1968) and Morrison (1970)
with the names used in this paper (Table 1). The American
Malacological Union List of Common and Scientific Names
[Turgeon et a/. (in press)] is incorporated as the basis for the
taxonomy used in this paper. However, the status of many
named subspecific varieties and ecophenotypes has not been
resolved. We list them here for clarity. Since the report by
Bickel (1968), almost half of the taxa have undergone tax-
onomic revision. Morrison (1970) and Johnson (1978) declared
Plagiola Rafinesque, 1819 available over Dysnomia Agassiz,
1852, but due to taxonomic questions about the type species,
we have chosen to use Epioblasma Rafinesque, 1831, the next
available generic name. Similarly, the change from Carun-
culina Simpson in Baker, 1898 to Toxolasma Rafinesque, 1831
involves five taxa (see Bogan and Parmalee, 1983). Additional-
ly, 12 taxa have been added to the state’s total list of species
while two, Fusconaia undata, (Barnes, 1823) and Amblema
peruviana (Lamarck, 1819) have been synonymized.

Bickel (1968) used 25 taxa originally described by
Rafinesque. Morrison (1970) included 26 nomenclatural
changes based on the priority of Rafinesque descriptions. In

STARNES AND BOGAN: MUSSELS OF TENNESSEE

Table 1. List of Tennessee unionids found in the Tennessee/Cumberland river systems.

Bickel (1968)

Actinonaias carinata (Barnes, 1823)
A. carinata gibba (Simpson, 1900)
A. pectorosa (Conrad, 1834)

Alasmidonta marginata Say, 1819

A. minor (Lea, 1845)

Amblema costata (Rafinesque, 1820)
A. costata perplicata (Conrad, 1841)
A. costata plicata (Say, 1817)

A. peruviana (Lamarck, 1819)
Anodonta grandis Say, 1829

A. grandis gigantea Lea, 1838

A. imbecillis Say, 1829

A. suborbiculata Say, 1831
Anodontoides ferussacianus (Lea, 1834)
Arcidens confragosus (Say, 1829)
Carunculina glans (Lea, 1831)

C. moesta (Lea, 1841)

C. moesta cylindrella (Lea, 1868)

C. parva (Barnes, 1823)

C. texasensis (Lea, 1857)

Conradilla caelata (Conrad, 1834)
Cumberlandia monodonta (Say, 1829)
Cyclonaias tuberculata (Rafinesque, 1820)
C. tuberculata granifera (Lea, 1838)
Cyprogenia irrorata (Lea, 1830)
Dromus dromas (Lea, 1834)

D. dromas caperatus (Lea, 1845)
Dysnomia arcaeformis (Lea, 1831)

. brevidens (Lea, 1831)

. capsaeformis (Lea, 1834)

. flexuosa (Rafinesque, 1820)

. florentina (Lea, 1857)

. florentina walkeri (Wilson and Clark, 1914)
haysiana (Lea, 1833)

. lenior (Lea, 1842)

. lewisi (Walker, 1910)

. stewardsoni (Lea, 1852)

SOvo00g0n0g00

D. torulosa (Rafinesque, 1820)

D. torulosa gubernaculum (Reeve, 1865)
D. torulosa propinqua (Lea, 1857)
D. triquetra (Rafinesque, 1820)
D. turgida (Lea, 1848)

Elliptio crassidens (Lamarck, 1819)

E. dilatatus (Rafinesque, 1820)

Fusconaia barnesiana barnesiana (Lea, 1838)
F. barnesiana bigbyensis (Lea, 1841)

F. barnesiana tumescens (Lea, 1845)

F. cuneolus cuneolus (Lea, 1840)

F. cuneolus appressa (Lea, 1871)

F. ebena (Lea, 1831)

F. edgariana (Lea, 1840)

F. edgariana analoga (Ortmann, 1918)

Morrison (1969)

Taxonomy used in this study

Actinonaias ligamentina (Lamarck, 1819)

Alasmidonta viridis (Rafinesque, 1820)

Toxolasma livida Rafinesque, 1831

Lemiox rimosus Rafinesque, 1831

Cyprogenia stegaria (Rafinesque, 1820)

Plagiola interrupta (Rafinesque, 1820)

Fusconaia pusilla (Rafinesque, 1820)

Actinonaias ligamentina

A. ligamentina gibba

A. pectorosa

Alasmidonta atropurpura (Raf., 1831)

A. marginata

A. raveneliana (Lea, 1834)

A. viridus

Amblema plicata (Say, 1817)

A. plicata perplicata

A. plicata plicata (Say, 1817)

A. plicata plicata

Anodonta grandis grandis

A. grandis corpulenta Cooper, 1834

A. grandis grandis

A. imbecillis

A. suborbiculata

Anodontoides ferussacianus

Arcidens confragosus

Toxolasma lividus glans

T. lividus lividus

T. lividus glans

T. cylindrella

T. parva

T. texasensis

Lemiox rimosus

Cumberlandia monodonta

Cyclonaias tuberculata tuberculata

C. tuberculata granifera

Cyprogenia stegaria

Dromus dromas dromas

D. dromas caperatus

Epioblasma arcaeformis

. biemarginata (Lea, 1857)

. brevidens

. capsaeformis

. flexuosa

. florentina florentina

. florentina walkeri

. haysiana

lenior

lewisi

. Stewardsoni

. obliquata (Raf., 1820)
(=sulcata Lea, 1829)

. torulosa

. torulosa cincinnatiensis (Lea, 1840)

. torulosa gubernaculum

. propinqua

. triquetra

. turgidula

Elliptio crassidens

E. dilatata

Fusconaia barnesiana

F. barnesiana bigbyensis

F. barnesiana tumescens

F. cuneolus

F. cuneolus appressa

F. ebena

F. cor cor (Conrad, 1834)

F. cor analoga

mMmmmmmmmmmm

mmmmmm

Pad

Table 1. (continued)

Bickel (1968)

F. flava (Rafinesque, 1820)

F. subrotunda (Lea, 1831)

F. subrotunda leseuriana (Lea, 1840)
F. subrotunda pilaris (Lea, 1840)

F. undata (Barnes, 1823)

Lampsilis anodontoides (Lea, 1831)

. anodontoides fallaciosa (Smith, 1899)
. fasciola Rafinesque, 1820

. orbiculata (Hildreth, 1828)

. ovata (Say, 1817)

. ovata satura (Lea, 1852)

. ovata ventricosa (Barnes, 1832)

mPererere

L. virescens (Lea, 1858)

Lasmigona complanata (Barnes, 1823)
L. costata (Rafinesque, 1820)

L. holstonia (Lea, 1838)

Lastena lata (Rafinesque, 1820)
Leptodea fragilis (Rafinesque, 1820)

L. leptodon (Rafinesque, 1820)
Lexingtonia dolabelloides (Lea, 1840)
L. dolabelloides conradi (Vanatta, 1915)
Ligumia recta latissima (Rafinesque, 1820)
L. subrostrata (Say, 1831)

Medionidus conradicus (Lea, 1834)
Megalonaias gigantea (Barnes, 1823)
Obliquaria reflexa (Rafinesque, 1820)
Obovaria olivaria (Rafinesque, 1820)

O. retusa (Lamarck, 1819)

O. subrotunda (Rafinesque, 1820)

O. subrotunda lens (Lea, 1831)

O. subrotunda levigata (Rafinesque, 1820)
Pegias fabula (Lea, 1838)

Plagiola lineolata (Rafinesque, 1820)
Plethobasus cooperianus (Lea, 1834)

P. cyphyus (Rafinesque, 1820)

P. cyphyus compertus (Frierson, 1911)
Pleurobema aldrichianum Goodrich, 1931
P. clava (Lamarck, 1819)

P coccineum (Conrad, 1836)

P cordatum (Rafinesque, 1820)

P. oviforme (Conrad, 1834)

P. oviforme argenteum (Lea, 1841)
P. oviforme holstonense (Lea, 1840)
P pyramidatum (Lea, 1831)

Proptera alata (Say, 1817)
P laevissima (Lea, 1830)

Ptychobranchus fasciolare (Rafinesque, 1820)

P. subtentum (Say, 1825)
Quaarula cylindrica (Say, 1817)
Q. cylindrica strigillata (Wright, 1898)

Q. intermedia (Conrad, 1836)
Q. metanevra (Rafinesque, 1820)

Q. pustulosa (Lea, 1831)
Q. quadrula (Rafinesque, 1820)

AMER. MALAC. BULL. 6(1) (1988)

Morrison (1969)

F. polita Say, 1834

F. polita lesueriana

F. polita pilaris

F. lateralis (Rafinesque, 1820)
Lampsilis teres (Rafinesque, 1820)

L. abrupta Say, 1831

L. cardium cardium (Raf., 1820)

Lasmigona badia (Rafinesque, 1831)

Megalonaias nervosa (Rafinesque, 1820)

Plethobasus Striatus (Rafinesque, 1820)
P. pachosteus (Raf., 1820)

Pleurobema obliguum Lamarck, 1819

P. obliquata Rafinesque, 1820
P permorsa Rafinesque, 1831
Potamilus alatus

P ohioensis (Rafinesque, 1820)

Quadrula bullata (Rafinesque, 1820)

Taxonomy used in this study

F. flava

F. subrotunda subrotunda
F. subrotunda lesueriana
F. subrotunda pilaris

F. flava

Lampilis teres anodontoides
. teres teres

. fasciola

abrupta

ovata

cardium satura

. cardium cardium

. Siliquoida (Barnes, 1823)
. virescens

Lasmigona complanata

L. costata

L. holstonia

Hemistena lata

Leptodea fragilis

L. leptodon

Lexingtonia dolabelloides
L. dolabelloides conradi
Ligumia recta latissima

L. subrostrata
Medionidus conradicus
Megalonaias nervosa
Obliquaria reflexa
Obovaria olivaria

O. retusa

O. subrotunda

O. subrotunda lens

O. subrotunda levigata
Pegias fabula

Ellipsaria lineolata
Plethobasus cooperianus
P cicatricosus (Say, 1829)
P. cyphyus

P. cyphyus compertus
Pleurobema aldrichianum
P. clava catillus

P coccineum

P cordatum

P. gibberum

P. oviforme

P. oviforme argenteum

P. oviforme holstonense

P rubrum (Rafinesque, 1820)
P plenum (Lea, 1840)
Potamilus alatus

P ohioensis (Rafinesque, 1820)
Ptychobranchus fasciolare
P. subtentum

Quaadrula cylindrica

Q. cylindrica strigillata

Q. fragosa (Conrad, 1835)
Q. intermedia

Q. metanevra

Q. nodulata (Rafinesque, 1820)
Q. pustulosa

Q. quadrula

Q. sparsa (Lea, 1841)

Perr rrrrer

STARNES AND BOGAN: MUSSELS OF TENNESSEE 23

Table 1. (continued)

Bickel (1968) Morrison (1969)

Taxonomy used in this study

Simpsoniconcha ambigua (Say, 1825)
Strophitus rugosus (Swainson, 1822)
Tritogonia verrucosa (Rafinesque, 1820)
Truncilla donaciformis (Lea, 1828)

T. truncata Rafinesque, 1820
Uniomerus tetralasmus (Say, 1831)
Villosa fabalis (Lea, 1831)

V. lienosa (Conrad, 1834)

V. nebulosa (Conrad, 1834)

V. picta (Lea, 1834)

Villosa teneltus (Rafinesque, 1831)

V. taeniata (Conrad, 1834)

V. trabalis (Conrad, 1834)

V. trabalis perpurpurea (Lea, 1861)
V. vanuxemensis (Lea, 1838)

Truncilla_ vermiculata (Rafinesque, 1820)

Simpsonaias ambigua
Strophitus undulatus (Say, 1817)
Tritogonia verrucosa

Truncilla donaciformis

T. truncata

Uniomerus tetralasmus

Villosa fabalis

V. lienosa

V. iris (Lea, 1830

V. taeniata picta (Lea, 1834)

V. taeniata punctata (Lea, 1865)
V. taeniata taeniata

V. trabalis

V. perpurpurea

V. vanuxemensis

this analysis, we have included three additional Rafinesque
species. Use of taxa originally described by Rafinesque is
perceived as controversial due to their convoluted
nomenclatural history (Bogan, Williams and Starnes, unpub.
data).

FACTORS AFFECTING DISTRIBUTION OF
UNIONIDS BY RIVER SYSTEM

MISSISSIPPI RIVER

The nature and size of the Mississippi River along the
western border of Tennessee virtually precludes a diverse
mollusk fauna. The river elevation annually fluctuates an
average of 6 m between winter highs and summer lows. The
substratum in shoal areas is sand and gravel while in pools
it consists of shifting sand and mud. With few species record-
ed from the Mississippi River proper, most have come from
oxbow lakes or tributary confluences.

Mississippi River tributaries in west Tennessee, with
migratory fishes providing the mechanism for dispersal, would
be expected to be relatively speciose. Unfortunately, agri-
cultural development of deep soils formed in loess and the
resulting deposition of sediments led to channelization of
these tributary rivers (Forked Deer, Obion, Wolf and
Loosahatchie) prior to documentation of their mussel fauna.

The Hatchie River (Table 2) appears to contain the on-
ly extant unionid fauna in Mississippi River tributaries in Ten-
nessee. Due to its relatively uniform sand/silt substratum,
diversity is relatively low in the Hatchie River. This limitation
of habitat diversity is typical of direct Mississippi River
tributaries. Most species recorded in the Hatchie River (D.
Manning, pers. comm.) occur in the Tennessee and Cumber-
land rivers; six species are new to the state list: Plectomerus
dombeyanus (Valenciennes, 1833), Uniomerus declivis (Say,
1831), Toxolasma texasensis (Lea, 1857), Obovaria jacksoniana
(Frierson, 1912), Potamilus purpurata (Lamarck, 1819) and
Villosa vibex (Conrad, 1834). Species such as Plectomerus
dombeyanus are widespread in Gulf Coast streams.

TENNESSEE RIVER

A total of 126 mussel taxa occur in the Tennessee River
and its tributaries. The Tennessee River, encompassing a
watershed of over 105,000 km?2, has been divided into
upper tributaries (Table 3) and middle and lower tributaries
(Table 4).

The French Broad and Holston rivers join to form the
Tennessee River. The Clinch and Powell rivers, originating in
the Ridge and Valley Province in southwestern Virginia, flow
into the Tennessee River. The underlying geology is folded
and faulted Paleozoic limestone lying in parallel northeast-
southwest ridges. Stream substrata are gravel, rubble and
bedrock of primarily limestone (Fenneman, 1938). Water is
hard and there are abundant nutrients [USEPA (United States
Environmental Protection Agency) STORET Database]. The
45 taxa that Ortmann (1924) considered ‘“‘Cumberlandian’’
have been recorded in this physiographic province.

The eastern headwater tributaries of the Tennessee
River arise in the Blue Ridge Province. The Watauga,
Nolichucky, French Broad, Pigeon, Little, Little Tennessee and
Hiwassee rivers originate along the western crest of the Blue
Ridge (600-800 m). Except in lower reaches, streams are
precipitous with soft water and low amounts of nutrients.
Geologically, the area is comprised of metamorphosed sedi-
mentary rocks, gneisses and schists (Fenneman, 1938).
Boulders, cobbles and siliceous rocks are typical substrata.
While there are endemic fish species such as brook trout
[Sa/velinus fontinalis (Mitchill)] in the Blue Ridge Province,
‘““Cumberlandian”’ unionid species are rare or totally absent.
Molluscan diversity and density, with few exceptions, in-
creases after these streams enter the Ridge and Valley Pro-
vince, lose gradient and change water chemistry (Bogan and
Starnes, 1983).

The Emory River (Table 3), a tributary to the lower
Clinch River, is a major stream draining the eastern portion
of the Cumberland Plateau. The Emory River crosses
geological strata that are characterized by Pennsylvanian
sandstone, shale and coal. The substratum is sandy with

24 AMER. MALAC. BULL. 6(1) (1988)

Table 2. List of Tennessee unionids found in the Mississippi River tributaries in Tennessee (N = Post 1960; R =

Prior to 1960).

North Fork
Species

Amblema plicata R R
A. plicata plicata R

Anodonta grandis

A. grandis corpulenta

A. imbecillis

A. suborbiculata
Arcidens confragosus
Elliptio crassidens
Fusconaia ebena

F. flava

F. flava trigona

Lampsilis cardium satura
L. siliquoidea N
L. teres teres R
L. teres anodontoides

Lasmigona complanata R
Leptodea fragilis

Ligumia subrostrata

Megalonaias nervosa R
Obovaria jacksoniana

Plectomerus dombeyanus R R
Plethobasus cyphus

Pleurobema cordatum

Potamilus ohiensis

P. purpurata

Quadrula pustulosa R
Q. pustulosa mortoni R

Q. quaarula R R
Strophitus undulatus

Toxolasma parva R
T. texasensis R
Tritigonia verrucosa R

Truncilla truncata R R
Uniomerus declivis

U. tetralasmus

Villosa lienosa

V. vibex

D
DDVUVUDD

DDVUDD

DDD

TOTAL TAXA 13 16

boulders, bedrock and shale. The water is soft, slightly acidic
and nutrient limited. A total of 22 taxa, including 11 Cumber-
landian endemics, have been recorded in this drainage, but
most occur in the lower reaches when the river enters the
Ridge and Valley Province and where the gradient has
decreased. The Sequatchie River, a southward flowing
tributary of the Tennessee River, drains the Southern
Cumberland Plateau. Twenty unionid species are listed from
the Sequatchie River (Table 4).

The Highland Rim Province dominates middle Ten-
nessee and encompasses several major tributaries of the
Tennessee River. Tributaries draining the crest of the Highland
Rim from the south, elevations of 250-300 m, include the Elk,
Flint and the Paint Rock rivers (the latter two do not contribute
taxa to the Tennessee fauna). The Buffalo River drains the

Reelfoot
Obion River Lake

Loosa-
Hatchie hatchie Wolf Horn
River River River Lake

N R
N
N
N
N R
N

R
N
N
N
N R R
N
N
N R
N
N
N
N
N
N
N
N R R
N R

R

N R
N N
N
N
N R R
N
N
N
N
N
32 7 6 1

interior of the southwestern Highland Rim while the Duck
River drains the eastern and western rim as well as the
southern Nashville Basin. These rivers are moderate in gra-
dient, nutrient enriched and have hard water. Substrata con-
sist of loose gravel or chert with limestone bedrock. Typical-
ly, these rivers are speciose with the Duck River (Table 4) hav-
ing 69 taxa; 25 Cumberlandian species inhabit the upper Duck
River. The Elk River (Table 4) similarly has 61 taxa recorded
from its waters. The Buffalo River (Table 4), a tributary to the
Duck River, is problematic; historically 27 taxa have been
recorded from this river (van der Schalie, 1973) but few species
have been recently collected in the drainage (Ahlstedt, 1986).
This is despite the fact that water quality appears acceptable
and faunal exchange could have occurred with the Tennessee
or Duck rivers since the substratum appears very similar to

STARNES AND BOGAN: MUSSELS OF TENNESSEE 25

Table 3. Mollusks of the Upper Tennessee River and its headwater tributaries (N = Post 1960; R = Prior to 1960; A = Archaeological).

French

Clinch Emory Watauga Broad Holston Little © Nolichucky Powell Tenn.
Species River River River River River River River River River
Actionaias ligamentina RN N N N
A. ligamentina gibba RNA R RN RN RN R
A. pectorosa RN R R R R RN R
Alasmidonta ravenelina N
A. marginata RN R RN N RN R
A. viridus R R R R
Amblema plicata RNA R R RN R RN RN R
Anodonta grandis grandis N
A. grandis corpulenta R
A. suborbiculata N
Cumberlandia monodonta RN R R RN R R
Cyclonaias tuberculata tuberculata RNA R RN RN N R
Cyprogenia stegaria RNA R R R
Dromus dromas dromas NA R N R
D. dromas caperatus R R RN R
Ellipsaria lineolata R R
Elliptio crassidens RNA R R RN RN RN R
E. dilatata RNA RN R R RN N RN RN R
E. dilatata subgibbosus R
Epioblasma arcaeformis RA R R R
E. biemarginata R R
E. brevidens RN R R R
E. capsaeformis RNA R R N RN RN R
E. fiorentina RA R
E. florentina walkeri R
E. haysiana RA R R R R
E. lenior R R R
E. lewisi R R R R
E. obliquata A
E. propinqua RA R R
E. stewardsoni RA R R
E. torulosa R
E. torulosa gubernaculum RNA R R R
E. triquetra RNA R R RN RN R
E. turgidula R R R R
Fusconaia barnesiana RNA R R RN N RN R
F. barnesiana bigbyensis RN R R R R R R
F. barnesiana tumescens R R R R R
F. cor analoga R R RN
F. cor RN N R
F. cuneolus appressa R R RN R R
F. cuneolus cuneolus NR R R RN R
F. subrotunda RN R R N RN
F. subrotunda lesuerianus RN R R R R R
F. subrotunda pilaris R R R R
Hemistena lata RN R N R
Lampsilis abrupta RNA R R
L. cardium R R R R RN R R
L. fasciola RNA R R R RN RN N RN R
L. ovata RNA N RN N RN R
L. virescens R R
Lasmigona complanata N
L. costata RN R R R R N N RN R
L. holstonia R R R R R R R R
Lemiox rimosus RNA R RN R
Leptodea fragilis RN N RN R RN R
L. leptodon R R R
Lexingtonia dolabelloides RNA R N R

26 AMER. MALAC. BULL. 6(1) (1988)

Table 3. (continued)

Clinch Emory Watauga
Species River River River
L. dolabelloides conradi R
Ligumia recta RNA
L. recta latissima RN
Medionidus conradicus RN R R
Obliquaria reflexa R
Obovaria retusa R
O. subrotunda subrotunda AR
O. subrotunda lavigata
Pegias fabula
Plethobasus cicatricosus A
P cooperianus RA
P. cyphyus RNA
P. cyphyus compertus
Pleurobema catillus R
P clava A
P coccineum R
P cordatum RNA
P. oviforme RN R
P. oviforme argenteum R R
P. oviforme holstonse R R
P plenum RNA
P rubrum RNA
Potamilus alatus RN N
Ptychobranchus fasciolare RNA RN
P subtentum RNA
Quadrula cylindrica cylindrica RNA
A. cylindrica strigulata R
Q. intermedia RA
Q. metanevra RNA
Q. pustulosa RNA N
Q. sparsa AN
Strophitus undulatus RN RN
Toxolasma cylindrellus
T. lividus glans R RN
T. lividus lividus R R
T. parva R
Truncilla truncta RN
Villosa fabalis R
V. iris RN R R
V. trabalis RA
V. perpurpurea RN R
V. vanuxemensis RNA R R
TOTAL TAXA 88 22 15

those rivers.

In addition to geology/water quality apparently affec-
ting mussel diversity and abundance, there is a strong cor-
relation between river drainage size and the occurrence of
mussels. In the Tennessee River, the smallest tributary to have
a diverse mussel fauna was Copper Creek (in Virginia) with
344.5 km2 of watershed (Ahlstedt, 1982). Other streams with
mussels had over 77.2 km? in drainage area.

SUMMARY OF TENNESSEE RIVER

The Tennessee River and its tributaries dominate the
state. A total of 126 mussel taxa has been reported from the

French
Broad Holston Little | Nolichucky Powell Tenn.
River River River River River River
R
R RN RN RN R
R
R RN RN R
R R
R R
R R
R
R R
R
R R R
R RN RN R
R R
R
R
R RN N R
R R RN R
R A RN R
R R R
R R R
R R R
R RN RN RN R
R R N RN R
R RN R
R RN R
R R
R R N R
R R
R RN RN RN R
R N
R R RN R
R
R
R R R R
R N R
R R R R
R R RN R RN R
R
R R RN RN RN R
40 79 20 30 48 63

Tennessee River drainage. This diversity is related to the
geology of the area where the headwater tributaries of the
river originate. The limestone enriched provinces of the head-
water drainages provide an ideal scenario for an expanded
mussel fauna: habitat diversity, abundant nutrients and
calcium enriched (hard) water. Due to man-induced habitat
changes (e.g. pollution and impoundments), the extant fauna
in the State is largely restricted to four Tennessee River
tributaries (i.e. the Duck, Elk, Clinch and Powell rivers). Con-
struction of the Columbia Reservoir on the Duck River began
in 1973 but was essentially halted in 1977. If that impound-
ment is completed, available habitat for Cumberlandian

STARNES AND BOGAN: MUSSELS OF TENNESSEE 27

mussel species will be further restricted by 32-48 km.

CUMBERLAND RIVER

The Cumberland River (Fig. 1) originates in the
Cumberland Mountain subprovince of the Cumberland
Plateau in southeastern Kentucky. It extends 1,105 km and
has a drainage of 48,000 km2. The Cumberland Plateau is
underlain by Pennsylvanian strata consisting of alternating
layers of shale, sandstone and coal. Water is soft and low in
dissolved nutrients. While the upper Cumberland River is con-
fined to Kentucky, the Big South Fork of the Cumberland River,
a major tributary, drains the western Cumberland Plateau in
Tennessee. Tributaries to the upper Cumberland River (Little
South Fork of the Cumberland, Rockcastle and Laurel rivers)
flow through Pennsylvanian-age strata through most of their
drainage. The Big South Fork has eroded through Pennsyl-
vanian into Mississippian strata (limestone). Twenty-five
unionid species have been recorded from the Big South Fork
drainage in Tennessee (Table 5).

As the Cumberland River enters Tennessee from Ken-
tucky it is joined by the Wolf, Obey and Roaring rivers. These
drain the eastern Highland Rim and possess substrata and
water chemistry similar to the Duck and Buffalo rivers. The
Obey River has 30 unionid species while the Roaring River
(Table 5) has 7 species.

As the Cumberland River enters the Nashville Basin,
it has reduced gradient and meanders westward across the
Basin until it re-enters the western Highland Rim. From the
south, the Cumberland River receives drainage from the
Caney Fork River (Southeastern Highland Rim) as well as the
Stones River (central Nashville Basin) (Schmidt, 1982). The
fauna of the Caney Fork (Table 5) is substantially reduced due
to a waterfall below the confluence of the Collins and Rocky
rivers. The Caney Fork River has 14 unionid taxa while the
Stones River (Table 5) has 49 taxa.

After re-entering the Highland Rim, the Cumberland
River flows westward through a deep alluvial floodplain. It
receives several major tributaries draining the surrounding
Highland Rim including the Harpeth and Red rivers and Yellow
Creek (Table 5). These tributaries have upland characteristics
with predominately chert-gravel substrata. The Harpeth and
Red rivers have 25 and 22 taxa, respectively (Table 5).

SUMMARY OF CUMBERLAND RIVER

A total of 85 mussel taxa has been recorded from the
Cumberland River and its tributaries in Tennessee. With 126
taxa recorded from the Tennessee River, this means that
numerous taxa including Cumberlandian species Quadrula
Sparsa (Lea, 1841), Lemiox rimosus Rafinesque, 1831 and Lex-
ingtonia dolabelloides (Lea, 1840) are absent from the
Cumberland River. All of the mussel species recorded from
the Cumberland River occur in the Tennessee River system.

The cause for this difference in total number of species
is probably related to geology. The Cumberland River head-
waters are in the nutrient-poor Pennsylvanian strata of the
Cumberland Plateau. These tributaries have relatively
depauperate faunas. It is only when streams cut through
Pennsylvanian strata into limestone that diversity increases

(Starnes and Bogan, 1982). A comparison of fauna in the Ten-
nessee and Cumberland rivers reveals that primarily the
headwater-mussel species are absent from the Cumberland
River. Thus, while these two rivers seem similar physio-
graphically, they are discretely different and this translates
into a slightly different mussel fauna.

CONASAUGA RIVER

This tributary to the Coosa River originates in the Blue
Ridge Province of northern Georgia and southern Tennessee.
The geology of the area is dominated by granite, gneisses,
schists and metamorphic rocks (Fenneman, 1938) that pro-
duce soft water with low nutrients. Mussels are absent from
this headwater area. After the river enters the Coosa Valley
(Ridge and Valley) Province, water becomes hard, nutrients
increase and bivalves begin to appear. The Conasauga River
in Tennessee contains 27 taxa (Table 6). Of these, Elliptio
dilatata (Rafinesque, 1820), Anodonta grandis corpulenta
Cooper, 1834, A. imbecillus Say, 1829, Lasmigona holstonia
(Lea, 1838), Toxolasma parva (Barnes, 1823), Medionidus con-
radicus (Lea, 1834), Villosa lienosa (Conrad, 1834) and V.
vanuxemensis (Lea, 1838) also occur in the Tennessee/Cum-
berland rivers and/or their tributaries. The remaining 19 taxa
are additions to the state species list and are typical of the
Coosa River system and Gulf coast streams (Table 6).

Near the Tennessee/Georgia border unionid species
diversity increases. An additional 15 species were collected
by Hurd (1974) immediately below that border but have not
been collected in Tennessee. These additional species may
be limited by habitat diversity or stream size from expanding
further upstream in the Conasauga River. Further research
into this area could be useful in understanding factors restric-
ting mussel distributions.

DISCUSSION

The earliest unionid faunal descriptions in Tennessee
were in the early 1800s. Subsequent malacological work has
tended to investigate the same rivers with diverse unionid
faunas while ignoring other major streams. It is ironic that no
comprehensive faunal surveys have been completed, until
recently, on the Conasauga, Hatchie or Mississippi rivers and
tributaries in Tennessee. Other works, such as ecological
studies of endemic species, are also very limited.

Since Ortmann’s work (1918, 1924, 1925) on the Ten-
nessee River system, rivers in this State have undergone con-
siderable change. There are now nine reservoirs on the main
Tennessee River, making it essentially a series of impound-
ments from its origin near Knoxville to its confluence with the
Ohio River. While the lack of complete historical data on the
early abundance and diversity of molluscan populations in
the Tennessee River (Table 6) and its tributaries confounds
any efforts to estimate the impact from man-made alterations,
changes have taken place. We can neither quantify the
change that has occurred in mussel populations during
historical times nor can we reliably predict what previous
changes portend for the health and survival of existing
populations.

28 AMER. MALAC. BULL. 6(1) (1988)

Table 4. Mollusks of the Middle and Lower Tennessee River and major tributaries (N = Post 1960; R = Prior to 1960; A = Archaeological).

Middle Tennessee River Lower Tennessee River
Little
Tenn. Hiwassee Sequatchie Tenn. Elk Duck Buffalo Tenn.

Species River River River River River River River River
Actionaias ligamentina A NR RN RN
A. ligamentina gibba NA RN RN
A. pectorosa R RNA RN R
Alasmidonta marginata N R RN R
A. viridus R R R RN R
Amblema plicata NA R RNA RN RNA RN
Anodonta grandis NA N NR RN N
A. grandis corpulenta N
A. imbecillis R RN
A. suborbiculata N
Arcidens confragosus N
Cumberlandia monodonta R N
Cyclonaias tuberculata NA R RNA RN RNA RN N
C. tuberculata granifera N RN
Cyprogenia stegaria A RNA R N
Dromus dromas A RNA NR
Ellipsaria lineolata RN NR R RN
Elliptio crassidens NA R R RNA N RN RN
E. dilatata RNA R RNA RNA RNA RN
Epioblasma arcaeformis A A
E. biemarginata R R
E. brevidens A A R RN
E. capsaeformis RA A RNA RNA
E. flexuosa A
E. florentina R A RN A
E. florentina walkeri R R
E. haysiana RA A R
E. lenior R
E. lewisi A
E. obliquata A
E. propinqua A A
E. stewardsoni A A
E. torulosa A RA RN R
E. triquetra A RN RNA
E. turgidula A R R
Fusconaia barnesiana RNA R R A RNA RNA RN
F. barnesiana bigbyensis R R R R R
F. barnesiana tumescens R R
F. cor RN
F. cuneolus RN
F. cuneolus appressa
F. ebena N RN
F. flava N
F. subrotunda RNA NA RN RN
Hemistena lata RN R R N
Lampsilis abrupta N N RN
L. cardium R R R R
L. fasciola RNA R RNA RNA RNA R
L. ovata RNA NA NA RNA N
L. teres anodontoides RN RN
L. teres teres N RN
Lasmigona complanata N N RN R
L. costata R RA RN RN R
L. holstonia R R R
Lemiox rimosus A A RN RNA
Leptodea fragilis N R N RN RN RN N

L. leptodon R

Table 4. (continued)

STARNES AND BOGAN: MUSSELS OF TENNESSEE

29

Middle Tennessee River Lower Tennessee River
Little
Tenn. Hiwassee Sequatchie Tenn. Elk Duck Buffalo Tenn.

Species River River River River River River River River
Lexingtonia dolabelloides NA RA RNA RNA N
L. dolabelloides conradi R R
Ligumia recta NA NA N
L. recta latissima N RN R
Medionidus conradicus NA RNA RNA
Megalonaias nervosa N RN RN RN
Obliquaria reflexa RN RN RN RN
Obovaria olivaria N RN
O. retusa A A R RN
O. subrotunda A A N RNA R
O. subrotunda lens R RN R R
Pegias fabula RA RA
Plethobasus cicatricosus A
P cooperianus A RA N RN
P. cyphyus NA NA RN
Pleurobema catillus R
P clava R A
P cordatum NA RNA N RN RN
P. oviforme RNA R RNA RNA R
P. oviforme holstonse R R R R
P. oviforme argenteum R R R R
P. plenum A RA R
P rubrum NA RNA NR R
P coccineum A R
Potamilus alatus NA R RNA N RN RN
P. ohioensis R N N
Ptychobranchus fasciolare A RNA RNA RN RN
P. subtentum A A RNA RA R
Quaarula cylindrica A R A RNA RNA
Q. fragosa R RN
Q. intermedia A RN RN
Q. metanevra NA RNA RN RN
Q. nodulata N
Q. pustulosa NA RNA N RN RN
Q. quadrula N RN RN
Q. sparsa R A
Strophitus undulatus N A RNA RNA R
Toxolasma cylindrellus RN R R R
T. lividus glans N RN
T. parva R
Tritigonia verrucosa R N RN RN RN
Truncilla donaciformis N N RN RN
T. truncata N RN R
Uniomerus tetralasmus N
Villosa fabalis N RN
V. iris R R R R RN RNA RN
V. taeniata RNA RNA RN
V. trabalis R
V. vanuxemensis RNA R A RNA RNA RN
TOTAL TAXA 50 12 20 66 61 68 27 45

ARCHAEOLOGICAL RECORD

The archaeological record is a valuable resource in
documenting the historical unionid fauna of Tennessee and
can provide clues to the early historical abundance and

distribution of mussel populations. It provides malacologists
with a significant supplement to historical mollusk collections.
The archaeological record can provide insight into the former
unionid fauna of what is now a dead or severely altered river

30 AMER. MALAC. BULL. 6(1) (1988)

(e.g. van der Schalie and Parmalee, 1960) or the past distribu-
tion of species not documented in historic collections (e.g. Par-
malee et a/., 1980).

Parmalee and Bogan (1986) discuss the late prehistoric
bivalve fauna of the lower Clinch River and document an ar-
chaeological assemblage richer and more diverse than that
reported by Ortmann (1918). The diverse prehistoric fauna of
the main channel of the Tennessee River in East Tennessee
has been alluded to by Parmalee (1966), Charles (1973) and
Bogan and Parmalee (1977). Parmalee et a/. (1982) document
the past unionid diversity of the Tennessee River above Chat-
tanooga, reporting 45 species from a series of archaeological
shell middens. They observed a major shift in the species
composition from late prehistoric samples to that fauna
represented in reaches impounded since the 1940’s. For ex-
ample, the most common species identified in these ar-
chaeological samples was Dromus dromas (Lea, 1834), an
endangered species (See Bogan and Parmalee, 1983) almost
extirpated from the main Tennessee River. The relative
dominance of Dromus in the prehistoric samples from the
Chickamauga Reservoir is comparable to those archaeolog-
ical assemblages from Widow’s Creek in northern Alabama
(Warren, 1975) and the large samples reported by Morrison
(1942) from the Pickwick Landing basin along the middle
stretch of Tennessee River in northwestern Alabama. The
relative abundance of the rest of the species is comparable
within the archaeological samples from the Clinch River,
Chickamauga Reservoir and the two Alabama studies. These
archaeological assemblages, when compared with the pre-
sent fauna, point to some major shifts in species assemblages
and abundance over the last 180 years. There has been
almost complete extirpation of all species of big river
Epioblasma sp. as well as other taxa such as Plethobasus
cooperianus (Lea, 1834), Actinonaias ligamentina (Lamarck,
1819), Quadrula intermedia (Conrad, 1836), Cyprogenia
stegaria (Rafinesque, 1820), Obovaria retusa (Lamarck, 1819)
and Pleurobema clava (Lamarck, 1819). These species have
been replaced by other taxa such as Ellipsaria lineolata
(Rafinesque, 1820), Obliquaria reflexa Rafinesque, 1820,
Tritogonia verrucosa (Rafinesque, 1820), Megalonaias nervosa
(Rafinesque, 1820) and Anodonta spp., which were essentially
absent from the archaeological record.

The naiad fauna of the Little Tennessee River, a
tributary of the Tennessee River in East Tennessee, was
surveyed and reported by Tennessee Valley Authority (1972)
as having a fauna of about 20 unionid species. Bogan (1982)
summarized the late prehistoric and early historic unionid
fauna of the Little Tennessee River as reported by Bogan
(1978, 1980, 1983), Robison (1978) and Bogan and Bogan
(1985), and had consisted of 46 species; an additional 14
species were expected but not found in the archaeological
samples. This reconstruction of the early historic fauna com-
pares favorably with other documented historic naiad faunas
from the Clinch, Holston and/or Powell rivers (Ortmann, 1918).

Archaeological bivalves recovered from the Eva site
on the west bank of the Tennessee River downstream from
the mouth of the Duck River document the former occurrence
of at least some of the ‘‘Cumberlandian’’ species as far

downstream as the mouth of the Duck River. Casey (1986)
documented the prehistoric occurrence of two Cumberlan-
dian species [Epioblasma arcaeformis (Lea, 1831), Dromus
dromas] near the mouth of the Tennessee River (River Mile
17.4) and the Cumberland River (River Mile 26) in Kentucky.
Parmalee (1982) and Parmalee and Klippel (1986) reported
the former occurrence of at least 26 species in the Duck River
based on a sample of naiads recovered from early and mid-
Holocene deposits. Robison (1986) included a discussion of
aborginal unionid samples from the Duck and upper Elk rivers.

Ortmann (1926b), in discussing the unionid fauna of
the Green River in Kentucky, noted the absence of Epioblasma
torulosa (Rafinesque, 1820) from the Cumberland River (ex-
cluding a probably spurious record from Walker). However,
Parmalee et a/. (1980) compared the modern fauna of the
Cumberland River with archaeological samples and
documented the former occurrence of E. torulosa in the
Cumberland River and noted that it was a common species
in the prehistoric faunal assemblage. Casey (1986) recorded
specimens of the E. torulosa complex from these same sites.

These examples clearly exemplify the importance of
archaeological material to the study of prehistoric and early
historic unionid distributions. The archaeological record is an
important supplement to modern collections and provides a
historical perspective on some of the changes in the naiad
fauna that have occurred in the past 180 years.

FAUNAL EXCHANGES

Evidence of faunal exchange between the Tennessee
and Cumberland rivers and the Ozark Region is supported
by archaeological records showing a larger range for
“Cumberlandian’ species than envisioned by Ortmann. Ort-
mann (1925) recognized that these two regions shared cer-
tain species, but did not elaborate. Cumberlandia monodon-
ta (Say, 1829) and Epioblasma turgidula (Lea, 1848) are shared
exclusively by these two regions. There is additional evidence
of faunal affinities with closely related taxa [i.e. Fusconaia
barnesiana (Lea, 1838) in the Tennessee and Cumberland
rivers and F. ozarkensis (Call, 1887) in the Ozarks]. Similar
affinities exist for Ptychobranchus fasciolare (Rafinesque,
1820) and P occidentalis (Conrad, 1836), and Cyprogenia
stegaria (Rafinesque, 1820) and C. alberti (Conrad, 1850). The
Tennessee and Cumberland river drainages share many
upland fish species groups and subgenera with the Ozarkian
region [for example: Notropis galacturus (Cope), N. telescopus
(Cope), Typhlichthys subterraneus Girard and Fundulus
catenatus (Storer) are exclusively shared by these regions
(Starnes and Etnier, 1986)]. These two regions exclusively
share fish and mussel species and yet these same species
are absent from adjacent tributaries to the Mississippi or Ohio
rivers.

Thus far, discussions of the Tennessee and Cumber-
land rivers have indicated that their mussel faunas are very
similar. Ortmann (1925) reported 10 taxa that were known to
be present in the Tennessee River but absent from the
Cumberland River. Of the Cumberlandian species found in
the Tennessee and Cumberland rivers, the following are ab-
sent from the lower Tennessee (Ortmann, 1924): Quadrula

STARNES AND BOGAN: MUSSELS OF TENNESSEE

Table 5. Species of the Cumberland River and its tributaries (N = Post 1960; R = Prior to 1960; A = Archaeological).

Cumber- Big So. Fork Caney
land Cumber- Obey Fork Stones Harpeth Red Roaring

Species River land River River River River River River River
Actinonaias ligamentina RNA R N R
A. ligamentina gibba R R R
A. pectorosa N R R N R
Alasmidonta atropurpurea N N
A. marginata R NR N
A. viridis N N N
Amblema plicata NA R RN
A. plicata perplicata R R
A. plicata plicata N
Anodonta grandis RN RN
A. imbecillis RN RN
Anodontoides ferussacianus R R
Cumberlandia monodonta RN R N
Cyclonaias tuberculata RNA R N R R
C. tuberculata granifera R
Cyprogenia stegaria RNA
Dromus dromas RNA R
Ellipsaria lineolata RN N
Elliptio crassidens RNA N R R
E. dilatata RNA N R N R R
Epioblasma arcaeformis A R
E. brevidens NA N R N
E. capsaeformis RA ? R R
E. flexuosa A
E. florentina RA R R R R
E. florentina walkeri RN 2 RN R R
E. havsiana RA
E. lenior RN
E. obliquata N R R
E. stewardsoni A
E. torulosa NA
E. triquetra N R
Fusconaia ebena RN
F. flava RNA RN R
F. subrotunda RNA R
Hemistena lata R R
Lampsilis abrupta RNA R
L. cardium R N RN R
L. fasciola RA N R RN R R R
L. ovata RNA N R R N R
L. teres anodontoides RN R N R R
L. teres teres RN
Lasmigona complanata RN R RN R R
L. costata RNA N R R RN R R R
Leptodea fragilis RN N
Lexingtonia dolabelloides NA
Ligumia recta latissima RNA N R N R
Medionidus conradicus N RN R
Megalonaias nervosa RN RN R
Obliquaria reflexa RNA R R N
Obovaria olivaria RN
O. retusa RNA
O. subrotunda RA R RN R RN
Pegias fabula N RN N
Plethobasus cicatricosus A R
P cyphyus RNA
P. cooperianus RNA

Pleurobema catillus R

32 AMER. MALAC. BULL. 6(1) (1988)

Table 5. (continued)

Cumber- Big So. Fork Caney
land Cumber- Obey Fork Stones Harpeth Red Roaring

Species River land River River River River River River River
P clava NA
P cordatum RNA N
P gibberum N
P. oviforme N R N
P. plenum RNA
P rubrum RNA N
P coccineum NA N N
Potamilus alatus RNA N R N R
P. ohioensis R R
Ptychobranchus fasciolare RNA N R N R
P. subtentum N R R R
Quadrula cylindrica RNA R N
Q. fragosa RN R
Q. metanevra RNA R
Q. pustulosa RNA N N R
Q. quaadrula N N
Simpsonaias ambigua N
Strophitus undulatus R N R N R R
Toxolasma lividus glans R
T. lividus lividus RN 2
T. parva N
Tritogonia verrucosa RN N R N R R
Truncilla donaciformis R N R
T. truncata RN R N R
Villosa iris A N R N
V. lienosa R N
V. taeniata picta R R
V. taeniata punctata R
V. taeniata RNA N R N RN
V. trabalis N RN
V. vanuxemensis N ? R
TOTAL TAXA 68 25 30 14 49 25 22 7

cylindrica strigillata (Wright, 1898); Plethobasus cyphus com-
pertus (Frierson, 1911); Alasmidonta raveneliana (Lea, 1834);
Villosa perpurpurea (Lea, 1861); Epioblasma torulosa guber-
naculum (Reeve, 1865); E. stewardsoni (Lea, 1852); E. lewisi
(Walker, 1910). A total of 87 mussel taxa have been reported
from the Cumberland River drainage while 126 taxa have been
recorded from the Tennessee River drainage. Thus, while
many species are shared, the fauna from the Cumberland
River does not include every species present in the Tennessee
River.

Faunal similarities occur between the two rivers
because of habitat and geological similarities instead of faunal
exchanges that would tend to make the faunas identical in
at least those rivers/streams where the exchange occurred
(see Starnes and Etnier, 1986). There are geological dif-
ferences between the two river drainages. Among these, there
is less physiographic diversity in the Cumberland River
drainage with the tributaries originating in Pennsylvanian
strata while those of the Tennessee River originate in Ridge
and Valley strata. This geologic dissimilarity between the Ten-
nessee and Cumberland tributaries probably contributes to

the dissimilarity in the total number of species. The Clinch
River, a part of the upper Tennessee River system, has had
89 taxa reported from its drainage. In contrast, the Stones
River, the tributary with the most diverse fauna in the
Cumberland River system, had only 49 taxa reported.

FAUNAL ALTERATIONS

As stated earlier, man-made river alterations have af-
fected mussel populations throughout recorded history. In im-
poundments the species Anodonta grandis Say, 1829; A. im-
becillis; A. suborbiculata Say, 1831; Obliquaria_reflexa;
Tritogonia verrucosa; Elliptio crassidens (Lamarck, 1819) and
Quadrula quadrula (Rafinesque, 1820) have expanded their
populations and distribution. While these species have pro-
liferated in reservoirs, those species requiring riverine en-
vironments for themselves or for their host fish species have
disappeared. Riverine species associated with the lower Ten-
nessee and Cumberland rivers appear least affected by im-
poundments, perhaps because there is little difference be-
tween a deep, slow-flowing river and a deep, slow-flowing
impoundment.

STARNES AND BOGAN: MUSSELS OF TENNESSEE

Table 6. Mollusks tabulated by river system (N = Post 1960; R = Prior to 1960; A = Archaeological).

Species

Actionaias ligamentina
A. ligamentina gibba

A. pectorosa
Alasmidonta atropurpurea
A. marginata

A. viridus

Amblema plicata

A. plicata perplicata

A. plicata plicata
Anodonta grandis

A. grandis corpulenta
A. imbecillus

A. suborbiculata
Anodontoides ferussacianus
Arcidens confragosus
Cumberlandia monodonta
Cyclonaias tuberculata
C. tuberculata granifera
Cyprogenia stegaria
Dromus dromas dromas
D. dromas caperatus
Ellipsaria lineolata
Elliptio arctata

E. crassidens

E. dilatata

E. dilatata subgibbosus
Epioblasma arcaeformis
biemarginata
brevidens
capsaeformis
flexuosa

florentina

florentina walkeri
haysiana

lenior

lewisi

metastriata

obliquata

propinqua
stewardsoni

. torulosa torulosa

. torulosa gubernaculum
. triquetra

. turgidula

mmmmmmmmmmmmmmmmm

Fusconaia barnesiana barnesiana

F. barnesiana bigbyensis
F. barnesiana tumescens
F. cor analoga

F. cor cor

F. cuneolus cuneolus

F. cuneolus appressa

F. ebena

F. flava

F. flava trigona

F. subrotunda

F. subrotunda lesuerianus
F. subrotunda pilaris

Tennessee River

Upper

RN
RN

RN

RN

Middle

RNA

RA

Lower

RN

Conasauga

River

Cumberland

River

Mississippi
River
Tributaries

RNA

RNA

RN
RN

33

34

Table 6. (continued)

AMER. MALAC. BULL. 6(1) (1988)

Tennessee River

Species Upper Middle Lower
Hemistena lata RN RN
Lampsilis abrupta RN N RN
L. altilis

L. cardium R R R
L. cardium satura

L. clarkiana

L. fasciola RN RNA RN
L. ornata

L. ovata RN RNA RN
L. siliquoidea

L. straminea claiborensis

L. teres RN
L. teres anodontoides

L. virescens R

Lasmigona complanata RN RN
L. costata RN RA RN
L. holstonia R R R
Lemiox rimosus RN A RN
Leptodea fragilis RN RN RN
L. leptodon RN R
Lexingtonia dolabelloides RN RA RN
L. dolabelloides conradi R R
Ligumia recta RN NA N
L. recta latissima RN N RN
L. subrostrata

Medionidus acutissimus

M. conradicus RN RN
Megalonaias nervosa N RN
Obliquaria reflexa R RN RN
Obovaria jacksoniana

O. olivaria N RN
O. retusa R A RN
O. subrotunda R A RN
O. subrotunda levigata R

O. subrotunda lens R RN
Pegias fabula R R
Plectomerus dombeyanus

Plethobasus cicatricosus A

P. cooperianus R RA RN
P. cyphyus RN NA RN
P._ cyphyus compertus RN NA RN
Pleurobema aldrichianum

P catillus R R
P clava RA

P. cordatum RN RNA RN
P georgianum

P. gibberum

P. hanleyanum

P johannis

P. oviforme RN R RN
P oviforme holstonse R R R
P. oviforme argenteum RA R
P perovatum

P. plenum RN AN R
P rubellum

P rubrum RN RA R
P coccineum R R R

P. troschelianum

Conasauga Cumberland

River River
R
RNA
N
RN
N
RNA
N
RNA
N
RN
RN
RNA
N
RN
NA
RNA
N
N RN
RN
RNA
RN
RNA
RNA
N
RA
RNA
RNA
RNA
N
R
NA
RNA
N
N
N
N
N
N
RNA
N
RNA
NA
N

Mississippi
River
Tributaries

RN

RN

RN

RN

RN

RN

STARNES AND BOGAN: MUSSELS OF TENNESSEE

Table 6. (continued)

35

Tennessee River

Lower

RN

Potamilus alatus

Species Upper Middle
RN RNA

P. ohiensis

P. purpurata

Ptychobranchus fasciolare RN RNA

P. greeni

P. subtentum RN A

Quaarula cylindrica RN RA

Q. cylindrica strigulata R

Q. fragosa

Q. intermedia RN A

Q. metanevra RN RNA

Q. nodulata

Q. pustulosa RN RNA

Q. pustulosa mortoni

Q. quaadrula

Q. sparsa N

Simpsonaias ambigua

Strophitus connasaugaensis

S. undulatus RN A

Toxolasma cylindrellus

T. lividus glans R

T. lividus lividus R

T. parva R R

T. texasensis

Tritigonia verrucosa RN

Truncilla donaciformis N

T. truncata RN

Uniomerus declivis

U. tetralasmus

Villosa fabalis R

V. iris RNR RN

V. lienosa

V. taeniata picta N?

V. taeniata punctata

V. taeniata taeniata

V. trabalis R R

V. trabilis perpurpurea RN

V. vanuxemensis RN R

V. vanuxemensis umbrans

V. vibex

TOTAL TAXA 94 73
ACKNOWLEDGMENTS

This compilation of published and unpublished information
could not have been accomplished without the assistance of John
B. Burch, University of Michigan; David H. Stansbery, Ohio State
University; Paul W. Parmalee, University of Tennessee; Don Manning,
McKenzie, Tennessee; Steven A. Ahlstedt, Tennessee Valley Authority
and Leroy Koch, State of Missouri. George M. Davis is acknowledged
for allowing us access to the Academy of Natural Sciences’
malacological collections. In addition we wish to acknowledge the
composition and typing assistance provided by Cynthia Bogan. Paul
W. Parmalee, James D. Williams and Robert Robertson reviewed and
constructively criticized drafts of this manuscript.

RN

RN

RN

RN
RN
RN

RN

RN

RNA

89

Mississippi
Conasauga Cumberland River
River River Tributaries
RNA
N RN
RN
RNA
N
R
RNA
RN
RNA
RNA RN
RN
N RN
N
N
RN RN
N R
RN
N N RN
RN
RN RN
RN
RN RN
N
N
N NA
N RN N
R
R
RNA
R
N RN
N
N N
27 85 35

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Date of manuscript acceptance: 20 January 1987

MORPHOLOGY OF GLOCHIDIA OF LAMPSILIS HIGGINSI
(BIVALVIA: UNIONIDAE) COMPARED WITH
THREE RELATED SPECIES

D. L. WALLER’, L. E. HOLLAND-BARTELS', AND L. G. MITCHELL?
'U. S. FISH AND WILDLIFE SERVICE
NATIONAL FISHERIES RESEARCH CENTER
P.O. BOX 818
LA CROSSE, WISCONSIN 54602-0818, U. S. A.
2IOWA STATE UNIVERSITY
DEPARTMENT OF ZOOLOGY
AMES, IOWA 50011, U. S. A.

ABSTRACT

Glochidia of the endangered unionid mussel Lampsilis higginsi (Lea) are morphologically similar
to those of several other species in the upper Mississippi River. Life history details, such as the timing
of reproduction and identity of host fish, can be readily studied if the glochidia of L. higginsi can be
distinguished from those of related species. We used light and scanning electron microscopy and
statistical analyses of three shell measurements, shell length, shell height, and hinge length, to com-
pare the glochidia of L. higginsi with those of L. radiata siliquoidea (Barnes), L. ventricosa (Barnes),
and Ligumia recta (Lamarck). Glochidia of L. higginsi were differentiated by scanning electron microscopy
on the basis of a combined examination of the position of the hinge ligament and the width of dorsal
ridges, but were indistinguishable by light microscope examination or by statistical analyses of
measurements. Analysis of variance and multivariate (principal component) analysis separated L. radiata
siliquoidea from the other three species by virtue of its larger size, but discriminant function analysis
classified only 38% of the glochidia of L. higginsi correctly compared with 83% of those of L. radiata

siliquoidea.

The glochidia of most unionid freshwater mussels are
obligate parasites on the gills or fins of fishes. Glochidia dis-
charged from the marsupial gills of females attach and en-
capsulate on fish and undergo organogenesis to the juvenile
stage (Coker et a/., 1921). Information on the life history and
recruitment of mussel species can be readily developed by
the collection and identification of glochidia. For example,
Zales and Neves (1982a, b) using light microscopy, determined
the timing of glochidial release, periods of infection, and the
identity of fish hosts for four lampsiline mussels by collecting
and identifying glochidia in stream drift and on fish gills.

Glochidia of the endangered Lampsilis higginsi (Lea)
are morphologically similar to those of several other species
of Lampsilinae in the upper Mississippi River (Surber, 1912,
1915). Before information about the reproductive cycle and
host fishes could be determined, a method for operational/field
identification of the glochidia of L. higginsi was required.

Several methods have been used to study glochidia.

Shell shape and gross features have been described by light
microscopy (Lefevre and Curtis, 1910; Surber, 1912, 1915; Ut-
terback, 1933; Inaba, 1941), shell dimensions have been
measured (Surber, 1912, 1915; Inaba, 1941; Wiles, 1975; Zale
and Neves, 1982a), and scanning electron microscopy has
been used by several researchers (Heffelfinger, 1969;
Calloway and Turner, 1978; Clarke, 1981, 1982; Rand and
Wiles, 1982). Although Surber (1912) provided camera lucida
drawings and measurements of glochidial length and width
from samples of Lampsilis higginsi, he provided no definitive
identification of the species. No further descriptions of
L. higginsi glochidia have been reported.

Our objective was to ascertain simple techniques that
could be used routinely in the field, including light microscope
examination and measurements of shell dimensions, to dif-
ferentiate the glochidia of Lampsilis higginsi from those of
three other lampsiline mussels (L. radiata siliquoidea
(Barnes), L. ventricosa (Barnes), and Ligumia recta (Lamarck)

American Malacological Bulletin, Vol. 6(1) (1988):39-43

39

40 AMER. MALAC. BULL. 6(1) (1988)

in the upper Mississippi River system. In addition, scanning
electron microscopy was used to study aspects of the com-
parative ultrastructure of the shells of these four species.

MATERIALS AND METHODS

Gravid females of 15 species of mussels, in addition
to Lampsilis higginsi, were collected from the upper Mississippi
River (Pools 7 and 10) by handpicking and brailing. After
preliminary examination, we selected L. radiata siliquoidea
(here termed L. radiata), L. ventricosa, and Ligumia recta for
detailed study because of the close similarity of their glochidia
to those of L. higginsi. We removed glochidia from live females
by using a hypodermic needle and syringe to flush the
marsupial portion of the gill. Glochidia that were infective and
therefore selected for examination responded by snapping
their valves shut when placed in a 1.0% NaCl solution. Other
glochidia came from females preserved in 10% formalin or
70% ethanol. In measuring length (maximum anterior-
posterior), height (maximum dorsal-ventral), and hinge length,
we examined 20 glochidia per female under a microscope
(100x) fitted with an ocular micrometer.

Data analyses were conducted using the Statistical
Analysis System (SAS Institute, 1979) at lowa State Universi-
ty, Ames. Statistical significance is defined as P <0.05.

Photographs were taken of representative specimens
of glochidia of each species for qualitative comparisons of
general shell features. All aspects of the shell were photo-
graphed, including lateral views showing the shape of the shell
and hinge, and the position, size, and shape of adductor mus-
cle, a dorsal view showing the hinge and beak sculpture, and
an anterior-posterior view showing the flange and shell gape.

Some glochidia of each species were fixed in 10% buf-
fered formalin and held in 70% ethanol for scanning electron
microscopy. Samples were prepared by critical point drying
and sputter coating with platinum palladium (Postek et a/.,
1980). Shells were studied at magnifications of 300x to
10,000x.

RESULTS AND DISCUSSION

STATISTICAL ANALYSIS

One-way analyses of variance revealed that overall
significant differences existed among glochidia of the four
species in the three morphometric characteristics measured.
However, the source of the difference was not due to Lamp-
silis higginsi, but to L. radiata which was significantly greater
in length, height, and hinge length than the other three
species (which did not differ significantly from one another).
(Table 1.) In addition, a multivariate (principal component)
analysis also did not separate L. higginsi from the other
species (Fig. 1). The first principal component had similar
loadings for all three characteristics (height = 0.59, length
= 0.60, hinge length = 0.54) and accounted for 77% of the
total variance in the correlation matrix. Component 2 (hinge
length = 0.83, height = 0.47, length = 0.29) accounted for

Plot of Principal | vs. Principal 2

Component 2

L. radiata

Component 1

L. ventricosa @ @
Ligumial

Fig. 1. Principal component analysis. The large size of Lampsilis
radiata glochidia separates it from the other forms along component 1.

only 16% of the variance. Again, L. radiata could be separated
from the other three species by its larger size, but L. higginsi
did not differ significantly from L. ventricosa and Ligumia recta.

Glochidia of Lampsilis higginsi were correctly classified
in 39% of the observations by discriminant analysis, but 55%
were misclassified as either L. ventricosa or Ligumia recta
(Table 2). Correct classifications were 50% for L. ventricosa
and 48% for Ligumia recta. Discriminant function analysis was
the most accurate for glochidia of L. radiata correctly classi-
fying 83% of the glochidia, 10% were misclassified as L.
higginsi.

LIGHT MICROSCOPY

The shape and appearance of shells of the four species
examined were so similar that identification by observation
with the light microscope was not possible (Fig. 2). Our
general observations of the shells were similar to those of
Lefevre and Curtis (1910) for hookless glochidia in shape of
the shell, the double margin around the periphery of the shell,
granulations on the lateral surface, the position and shape
of adductor muscle, and the presence of two pairs of micropro-
jections. When profiles of the shells of each species were com-
pared by overlaying transparencies of shells of the same size,
no obvious differences in shape could be detected, although
about 4% of the glochidia in one female Ligumia recta were
much higher than most glochidia of this species (height x =
296 nm). The relative position of the hinge ligament could be
discerned in some glochidia of each species at 40x. The hinge
ligament in L. recta was centrally located whereas that in the
three Lampsilis species was more posterior. The position of
the adductor muscle was not considered for use in identifica-
tion because the larval adductor muscle is lost soon after a
glochidium attaches to a fish. Other features of the glochidium
were not adequately resolved by light microscopy to be useful
for species identification.

SCANNING ELECTRON MICROSCOPY

Scanning electron microscopy showed all four species
have similar surface features. A series of semi-circular ridges

WALLER ET AL.: LAMPSILIS GLOCHIDIA 41

Fig. 2. Glochidia of Lampsilis higginsi: light microscope photograph (scale bar = 100 nm). Fig. 3. Scanning electron micrograph of characteristic
features of shell valve of Lampsilinae glochidia (scale bar = 100 um). Fig. 4. Scanning electron micrograph (anterior view) showing flattened
dorsal ridges (D) of Lampsilis higginsi (scale bar = 100 um). Fig. 5. Ventral flange (F) and a portion of the lateral shelf of glochidium of L.
ventricosa (scale bar = 20 um). Fig. 6. Ventral flange showing fine tooth-like projections and pits on internal surface (arrow) of glochidium
of L. ventricosa (scale bar = 10 um). Fig. 7. Internal view of the glochidium as seen in the gaping shell: mantle, adductor muscle, and micropro-
jections (arrow) (scale bar = 100 um). Fig. 8. Internal view of hinge ligament, placed slightly posterior in Lampsilis radiata (scale bar = 100
um). Fig. 9. External sculpturing of the shell at the dorsal edge in Lampsilis ventricosa glochidium (scale bar = 50 um).

42 AMER. MALAC. BULL. 6(1) (1988)

Table 1. Mean measurements (standard deviations in parentheses)
of glochidia of four Lampsilinae mussels. Means for each measured
characteristic with the same superscript are not significantly different
from each other (P = 0.05) (Student-Newman-Keul’s test of means).

Species Number Number Measurements (um)
of of ; :

glochidia females “eight Length Hinge

Lampsilis higginsi 96 3 2599 = 2152 = 1108
(8.0) (4.2) (4.2)

L. radiata 220 11 2715 + =228 = 420P
(1.2) (08) (05)

L. ventricosa 556 19 2572 = 2162 ~—- 1074
(0.9) (0.8) (0.5)

Ligumia recta 180 9 2599 =2138 = 1078

(0.9) (0.5) (0.5)

on the lateral surface become wrinkled near the hinge (Fig.
3). In addition, each valve has many pits on both the internal
and external surface, which have sometimes been interpreted
as pores (Arey, 1924; Zs.-Nagy and Labos, 1969; Calloway and
Turner, 1978; Rand and Wiles, 1982). When viewed from the
lateral external surface, the shell does not appear to be
porous, but in cross-sectional and internal examinations of
the valves, the pits appeared to be continuous. This apparent
discrepancy could have been explained by Calloway and
Turner (1978), who noted that the external surface appeared
perforated only at accelerating voltages of 20 kilovolts and
greater. They concluded that the periostracum was not per-
forated and that the appearance of pores on the outer sur-
face was an artifact of the scanning electron microscope.
Perhaps electrons penetrate the periostracum at 20 kilovolts
making it appear transparent and the pits appear as pores.
Posterior and anterior edges of each valve are flattened near
the dorsal aspect, forming a smooth surface about 65 um long
(dorso-ventral) and 25-40 um wide (medial-lateral) here re-
ferred to as dorsal ridges (Fig. 4). The peripheral edges of
the valves are turned inward to form a continuous shelf around
the inner margin. Ventrally, the shelf forms a flange or lip be-
lieved to be analogous to the hook of anodontine mussels
described by Lefevre and Curtis (1910) (Fig. 5). The flange

Table 2. Summary of percent of each species classified as either
Lampsilis higginsi, L. radiata, L. ventricosa, or Ligumia recta by
discriminant analysis.

Known Percentage classified into species
species Number
Lampsilis L. L. Ligumia_ of speci-
higginsi radiata ventricosa recta mens
Lampsilis
higginsi 39 6 22 33 96
L. radiata
siliquoidea 10 83 5 2 220
L. ventricosa 19 8 50 24 556
Ligumia recta 20 5 27 48 180

is about 13-19 ~m wide and extends the width of the ventral
shell margin. Fine, tooth-like projections, previously described
as microstyles (Clarke, 1985), cover all except the proximal
one-third of the flange (Fig. 6). The microstyles decrease in
length to micropoints on the inner edge of the flange. In all
four species, the microstyles are arranged in irregular ver-
tical rows and about 14-17 rows cover the flange from the in-
ner to the outer edge. The inner shell margin provides an at-
tachment site for the mantle, a thin sheet of tissue covering
the inner valve surface except in the region of the adductor
muscle. The single adductor muscle was also seen internal-
ly near the dorsal margin. A pair of cylindrical microprojec-
tions, about 24-26 um long, previously described as ‘‘sensory
hairs”’ (Lefevre and Curtis, 1910), is near the ventral margin
of the valve (Fig. 7). At the dorsal edge of the valve, the shelf
folds inward forming an articulating surface for junction of the
valves. The larval ligament connects the valves at this hinge
line (Fig. 8).

Table 3. Width of the dorsal ridge of the four Lampsilinae species.
Mean dorsal ridge widths with the same superscript are not
significantly different from each other (P = 0.05) (Student-Newman-
Keuls’ test of means).

Species N Width of ridge (um)
Mean SD Range
Lampsilis higginsi 9 27.202 1.75 25.0-29.2
L. radiata 9 3348 313 280-379
L. ventricosa 10 34.70 306 300-400
Ligumia recta 8 28.662 0.96 25.8-30.0

We concentrated on three features of the shell in our
efforts to distinguish among the species: (1) position of the
hinge ligament; (2) width of the flattened dorsal ridges; (3)
sculpturing on the lateral shell surface. The first two features
proved to be the most useful for separating Lampsilis higgin-
si. Hinge ligaments were central in glochidia of Ligumia rec-
ta, whereas they were slightly more posterior in L. higginsi,
L. ventricosa, and L. radiata. The dorsal ridges of each valve,
measured at their greatest width, differed significantly among
species (Table 3). The ridge width was usually narrower in
L. higginsi and Ligumia recta (250-300 »m) than in L. ven-
tricosa and L. radiata (280-400 um).

The shell sculpture showed no major differences
among the species, though there was some subtle variation.
We attempted to identify photographs of each species on the
basis of shell sculpture alone, but could not consistently detect
a representative pattern on each shell.

CONCLUSION

The objective of this study was to find an operational/
field method for routine identification of Lampsilis higginsi.
Light microscopy and statistical analyses of shell dimensions
were found to be inadequate for species differentiation. Scan-
ning electron microscopy can be used to differentiate glochidia

WALLER ET AL.: LAMPSILIS GLOCHIDIA 43

of L. higginsi from the other three species on the basis of the
position of the hinge ligament and the width of the dorsal
ridges, but the technique is expensive and impractical for iden-
tification of small samples of glochidia collected in the field.
The technique could be of use when there is justification for
a significant investment of time and expense in identification
of glochidia. Hoggarth and Cummings (pers. comm.) used
scanning electron microscopy to identify glochidia of Anodonta
grandis grandis Say on fish in field collections and suggested
this technique was more labor efficient than artificial infec-
tion experiments for determining host fishes. On the contrary,
we have found that artificial infection requires much less
equipment, training, and expense than scanning electron
microscopy and is more practical for routine use.

Laboratory culture of glochidia and juveniles (Isom and
Hudson, 1982; Hudson and Isom, 1984) could be another
route for developing early life histories. Investigators could
follow the growth of a mussel and document developmental
stages at which Lampsilis higginsi can be positively differen-
tiated from related species by light microscopy. One could
then verify fish hosts by holding field-collected fish in the
laboratory until juvenile mussels have dropped off and
developed into an identifiable stage. Recruitment of a species
could also be evaluated by identification of juveniles in the
field.

LITERATURE CITED

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Calloway, C. B. and R. D. Turner. 1978. New techniques for preparing
shells of bivalve larvae for examination with the scanning elec-
tron microscope. Bulletin of the American Malacological Union
for 1977:17-24.

Clarke, A. H. 1981. The tribe Alasmidontini (Unionidae: Anodontinae),
Part 1: Pegias, Alasmidonta, and Arcidens. Smithsonian Con-
tributions to Zoology 326:1-101.

Clarke, A. H. 1985. The tribe Alasmidontini (Unionidae: Anodontinae),
Part Z: Lasmigona and Simpsonaias. Smithsonian Contribu-
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freshwater mussels (Unionidae) in a laboratory setting. Nautilus
98:129-135.

Inaba, S. 1941. A preliminary note on the glochidia of Japanese
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29:14-23,

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freshwater mussel glochidia. Nautilus 96:147-135.

Lefevre, G. and W. C. Curtis. 1910. Reproduction and parasitism in
the Unionidae. Journal of Experimental Zoology 9:78-115.

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1980. Scanning Microscopy: a Student’s Handbook. Ladd
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Rand, T. G. and M. Wiles. 1982. Species differentiation of the glochidia
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1829 (Mollusca: Unionidae) by scanning electron microscopy.
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771. 13 pp.

Surber, T. 1915. Identification of the glochidia of freshwater mussels.
Bulletin of the United States Bureau of Fisheries Document No.
813. 10 pp.

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Date of manuscript acceptance: 3 August 1987

RESEARCH NOTE

A TECHNIQUE FOR TRAPPING SANDFLAT OCTOPUSES

JANET R. VOIGHT
DEPARTMENT OF ECOLOGY AND EVOLUTIONARY BIOLOGY
UNIVERSITY OF ARIZONA
TUCSON, ARIZONA 85721, U. S. A.

ABSTRACT

An intertidal population of Octopus digueti Perrier and Rochebrune was sampled without ap-
parent sex or size bias (except for the smallest size classes) by placing artificial shelters in the inter-
tidal zone. Comparisons of captures between the octopuses’ natural shelters, large gastropod shells,
and the artificial shelters, glass bottles, revealed no differences in the sex or size of the octopuses
captured. The bottle trap technique is an inexpensive means of sampling O. digueti. The technique
provides large numbers of untraumatized octopuses and can define the local species distribution in

this potentially shelter-limited population.

A basic problem in the study of octopus populations
is that of reliable sampling. Most workers have employed hand
capture by divers armed with chemical irritants (e. g. Smale
and Buchan, 1981; Ambrose, 1984; Hartwick et a/., 1984; Aron-
son, 1986), or capture by trawls (Mangold and Boletzky, 1973;
Hatanaka, 1979; Guerra, 1981; Boyle and Knobloch, 1982;
Boyle, 1986). Both techniques have inherent drawbacks.
Divers locate more large animals than small ones, and are
limited by water clarity, depth restrictions, past experiences
of individual divers, the type of shelter available to the oc-
topuses and the persistence of den middens (Ambrose, 1983;
Hartwick, 1983; Van Heukelem, 1983). Trawl captures are
limited to species occurring on trawlable bottoms, and are
biased by net mesh size and varying trawl times (Boyle, 1983).

Beginning in ancient times, a number of widely
separated fishing cultures have captured octopuses by plac-
ing artificial sheiters in the sea and recovering them after the
octopuses have taken up residence. Such trapping techniques
have been successful for Octopus dofleini (Wulker) in the
northeast Pacific, O. briareus Robson in the Caribbean, O.
tetricus Gould in Australia and O. vulgaris Cuvier in the
Mediterranean (Lane, 1957; Roper et a/., 1984).

Current uses of traps in the study of octopuses have
been limited to providing a few untraumatized octopuses for
laboratory studies (Nixon, 1969; Joll, 1976, 1977) and to
assessing the fisheries potential of a population (Whitaker and
DeLancey, 1986). Although octopuses use a wide variety of
shelters in the wild, selection experiments have revealed that
octopuses show an aversion to transparent shelter in both

laboratory (Mather, 1982) and field (Aronson, 1986) studies.
Shelters with narrow apertures are preferred by Octopus
joubini Robson (Mather, 1982).

This paper describes a trapping technique that has pro-
ven useful in the study of Octopus digueti Perrier and
Rochebrune, a small (generally less than 40 g) octopus oc-
curring on sandy bottoms throughout the Gulf of California.
This species typically uses the shelter provided by vacant
gastropod and bivalve shells (Hochberg, 1980) that can be
limiting, since individuals are often found under shell
fragments, in bottles or cans, or even buried in the sediment
(Perrier and Rochebrune, 1894; pers. obs.). This technique
uses brown glass bottles as artificial shelters that serve as
inexpensive and reliable traps. They provide a means of
sampling the population and can provide relatively un-
traumatized octopuses for laboratory studies.

MATERIALS AND METHODS

The study area was located in Choya Bay, Sonora,
Mexico. The bay is a 5 km2 area of sandflats, located about
5 km northwest of the town of Puerto Penasco, in the north-
ern Gulf of California. Extreme vertical tidal ranges (to 7 m)
and the gentle slope of the bottom made intertidal trapping
feasible. Octopus digueti is common in Choya Bay, especial-
ly in areas of permanent water cover such as tide pools or
channels where shell refuges are abundant.

Bottle traps used in this study were barrel-shaped,
325 ml brown glass beer bottles (Cerveza Corona) that taper

American Malacological Bulletin, Vol. 6(1) (1988):45-48

45

46 AMER. MALAC. BULL. 6(1) (1988)

to a 17 mm neck diameter. A nylon electrician’s cable tie
secured around the bottle neck and a metal paper clip slipped
through the cable tie fastened each trap to an anchor line
and facilitated easy removal. Shells of Muricanthus nigritus
Philippi and Hexaplex erythrostomus Swainson with apertures
ranging from 29x42 mm to 76x98 mm were used as controls
for the bottle traps, both for estimating capture rates, and for
sampling larger octopuses that do not utilize bottle traps. The
shells were assembled into trap lines using the same method
as the bottle traps, with a cable tie inserted through two holes
drilled in the outer whorl of each shell.

Traplines consisted of 10 traps attached to loops tied
at one meter intervals on 40 or 50 pound test (18 or 23 kg)
nylon monofilament. Each line was staked at both ends by
a 0.3 m length of steel reinforcing bar driven into the
substratum. Three lines of shell traps and nine lines of bottle
traps were set between 10 July and 24 Sept 1984. Most
traplines were staked in optimal habitat for the octopuses,
areas with abundant shell debris and with water during the
lowest tides. To determine the vertical distribution of the
species in the intertidal zone, lines were staked from —1.3 to
+0.7 m. Both the outer flat habitat, an area with coarse sand
and abundant shell debris, and the inner flat habitat, an area
of fine sediment and few shells (Flessa and Ekdale, 1987),
were sampled by bottle traps.

All traplines were left staked in the intertidal zone
throughout the duration of the study. They were checked at
24 hour intervals during spring tides when low tides were at
—0.6 m or lower. The number of traps containing octopuses,
and the number of traps lost were recorded at each inspection.

Traps with resident octopuses were removed from the
line and replaced with empty traps. Captured octopuses were
taken in their traps to the marine laboratory at the Centro de
Estudios de Desiertos y Oceanos (CEDO) near Puerto
Penasco and placed in aquaria. Each individual was induced
to leave its trap by draining the water. All octopuses were nar-
cotized by a brief immersion in a 3-4% ethanol-seawater solu-
tion. Body weight was determined on a triple beam balance,
after water was drained from the mantle. A variety of
measurements were also made on each individual, of which
head width is reported here. The hyaline cranium (Boyle et
al., 1986) is the most rigid part of the octopus body and, as
such, could be indicative of size selection imposed by the nar-
row neck of the bottle-traps. The sex of each individual over
15.0 g was determined by the presence in males of a hec-
tocotylized third right arm, and by its absence in females. Oc-
topuses under 15.0 g were considered to be juveniles. The
octopuses were returned to within 800 m of the trap locality
at the next suitable low tide.

RESULTS

Of 2,244 total traps set overnight for twenty-one nights,
317 captured octopuses, for an overall capture rate of 14.1%.
Traplines placed in optimal octopus habitats in the outerflats
routinely contained octopuses. However, traplines in the in-
nerflats never captured any octopuses. Captures were rare
where the outer and innerflats intergraded. In optimal habitats,

Table 1. Sexual composition of Octopus digueti sampled by bottle
traps and shell traps. Individuals weighing less than 15 g were con-
sidered juveniles and were excluded from this analysis. Chi-square
for deviation from 1:1 sex ratio for bottle trap sample y2=2.66, p > 0.05;
for shell trap sample y?=.38, p>0.05.

Bottle Traps Shell Traps
Males 88 23
Females 111 19
Juveniles 55 2

shell traps were statistically more effective than were bottle
traps (18.3% versus 11.7%, y2=6.85, p<.01). Trap losses from
breakage and dislodgement over the three month period were
18.8% for the shell traps and 26.7% for the bottle traps.

Potential competitors for shelter in the bottle traps were
not seen. However, juvenile spotted sand bass (Paralabrax
maculatofasciatus Steindachner) occasionally took refuge in
the shell traps and could have excluded the octopuses.

Sex ratios of adult Octopus digueti captured by both
types of trap were not significantly different from 50:50 (chi-
square analysis with a Yates correction factor, Table 1). Head
widths of animals captured by bottle traps were not significant-
ly different from those captured by shell traps (p>0.10,
Kolmogorov-Smirnov Two-sample Test), although the bottle
traps captured more small individuals (Table 2).

The mortality observed in this study was limited to two
animals that died as a result of wedging themselves into bot-
tle necks. Otherwise, captured octopuses survived the trip to
the laboratory and the narcotization.

Table 2. Number of head widths of individual Octopus digueti from
bottle traps and shell traps.

Head width in mm Bottle traps Shell traps
10.0-11.9 1 1
12.0-13.9 14 0
14.0-15.9 34 0
16.0-17.9 49 5
18.0-19.9 72 10
20.0-21.9 69 18
22.0-23.9 14 10
24.0-25.9 1 0

DISCUSSION

Bottle traps provided an inexpensive, reliable means of
collecting large numbers of Octopus digueti. The total capture
rate (14.1%) compares favorably with capture rates obtained
by snap-trapping small mammals (Voight and Glenn-Lewin,
1979), although during a one-year study of this O. digueti
population, total capture rates were strongly affected by
seawater temperatures (Voight, unpub. data). Whitaker and
DeLancy (1986) reported a 26% capture rate in a potting study
of O. vulgaris sampled at intervals of from several days to
several weeks along the Atlantic coast of North America. In
their study, as in this one, octopuses collected in traps were
spared injuries associated with trawl captures and the ex-

VOIGHT: OCTOPUS TRAP TECHNIQUE 47

posure to chemicals required for hand collection by divers,
hence they were relatively untraumatized.

The comparison of capture rate between shells and
bottles showed that shells were more effective as traps and
less likely to be lost. However, bottles had an advantage in that
they were more easily acquired than were large numbers of
suitable gastropod shells, and they had narrow apertures. In
the laboratory, Octopus joubini, a small sandflat octopus from
the Gulf of Mexico and the Caribbean, prefer shelters with
relatively narrow apertures to those with wide apertures
(Mather, 1982). A similar preference in O. digueti could ex-
plain the attractiveness of the narrow-necked bottles with slop-
ing sides as shelter. The funnel-shaped upper third of the bot-
tle allowed small individuals to contact a solid wall, if they
remained near the bottle neck. The barrel shape allowed large
individuals, once past the narrow aperture, ample space while
maintaining contact with the solid wall. Thus, the shape of
the bottle assured little size bias.

The aversion to shelters that allow light penetration,
reported in Octopus joubini and O. briareus (Mather, 1982;
Aronson, 1986), could have been minimized in this study by
the use of brown glass. This aversion, if present in O. cigueti,
could have reduced the capture rate of the bottle traps.

Very small individuals, less than 18 mm head width,
were underrepresented by both techniques. Since Octopus
digueti produces young in the study area that immediately
assume a benthic existence (Hanlon and Forsythe, 1985), it
is assumed that all sizes of octopuses were available for trap-
ping. Young octopuses are likely to be more secretive and
less mobile than are adults, which may explain their lower
capture rate.

No sex bias was apparent in Octopus digueti with either
trap technique in the present study (Table 1), the sexes are
thought to be equally represented in other Octopus popula-
tions (Wells and Wells, 1977; Guerra, 1981; Smale and
Buchan, 1981; Aronson, 1986). The strongly female biased
sex ratios that have been observed in O. dofleini have been
attributed to behavioral differences between the sexes (Hart-
wick et al., 1984). Field studies of Eledone cirrhosa (Lamarck)
also show a female biased sex ratio, which has been at-
tributed to female migration into shallow waters (Boyle, 1983).

In addition to monitoring the population, the bottle trap
technique effectively demonstrated the local species distribu-
tion. The capture rate of octopuses declined to zero with the
change in substratum from coarse sand and shells to fine
sand with few shells. Without the trap technique, extensive
surveys would have been required to define the upper limit
of the species’ range in the intertidal zone.

ACKNOWLEDGMENTS

| thank K. W. Flessa, P. A. Hastings, C. M. Lively, D. A. Thom-
son and R. B. Toll for comments on the manuscript, P. A. Hastings
and C. M. Lively for statistical advice, and the staff of CEDO for hous-
ing and laboratory facilities. This research was conducted under the
auspices of Mexican Permits #2999 and #0475 to J. R. Hendrickson
who was instrumental in the design and implementation of the
technique.

LITERATURE CITED

Ambrose, R. F. 1983. Midden formation by octopuses: the role of
biotic and abiotic factors. Marine Behavior and Physiology
10:137-144.

Ambrose, R. F. 1984. Food preferences, prey availability, and the
diet of Octopus bimaculatus Verrill. Journal of Experimental
Marine Biology and Ecology 77:29-44.

Aronson, R. B. 1986. Life history and den ecology of Octopus briareus
Robson in a marine lake. Journal of Experimental Marine
Biology and Ecology 95:37-56.

Boyle, P. R. 1983. Eledone cirrhosa. In: Cephalopod Life Cycles, Vol.
1, P. R. Boyle, ed. pp. 365-386. Academic Press, London.

Boyle, P. R. 1986. A descriptive ecology of Eledone cirrhosa
(Mollusca:Cephalopoda) in Scottish waters. Journal of the
Marine Biological Association of the United Kingdom
66:855-865.

Boyle, P. R. and D. Knobloch. 1982. On growth of the octopus
Eledone cirrhosa. Journal of the Marine Biological Association
of the United Kingdom 62:277-296.

Boyle, P. R., M. S. Grisley and G. Robertson. 1986. Crustacea in
the diet of Eledone cirrhosa (Mollusca:Cephalopoda) deter-
mined by serological methods. Journal of the Marine Biological
Association of the United Kingdom 66:867-879.

Flessa, K. W. and A. A. Ekdale. 1987. Paleoecology and taphonomy
of Recent to Pleistocene intertidal deposits, Gulf of Califor-
nia. In: Geological diversity of Arizona and its margins: Excur-
sions to choice areas, G. H. Davis and E. M. VandenDolder,
eds. pp. 295-308. Arizona Bureau of Geology and Mineral
Technology, Tucson. Special paper #5.

Guerra, A. 1981. Spatial distribution pattern of Octopus vulgaris. Jour-
nal of Zoology London 195:133-146.

Hartwick, B. 1983. Octopus dofleini. In: Cephalopod Life Cycles, Vol.
1, P. R. Boyle, ed. pp. 277-291. Academic Press, London.

Hartwick, E. B., R. F. Ambrose and S. M. C. Robinson. 1984.
Dynamics of shallow-water populations of Octopus dofleini.
Marine Biology 82:65-72.

Hanlon, R. T. and J. W. Forsythe. 1985. Advances in the laboratory
culture of octopuses for biomedical research. Laboratory
Animal Science 35:33-40.

Hatanaka, H. 1979. Spawning seasons of the common octopus off
the northwest coast of Africa. Bulletin of the Japanese Socie-
ty of Scientific Fisheries 45:805-810.

Hochberg, F. G. 1980. Class Cephalopoda. /n: Common intertidal
invertebrates of the Guif of California, R. C. Brusca, ed. pp.
201-204. University of Arizona Press, Tucson.

Joll, L. M. 1976. Mating, egg-laying and hatching of Octopus tetricus
(Mollusca:Cephalopoda) in the laboratory. Marine Biology
36:327-333.

Joll, L. M. 1977. Growth and food intake of Octopus tetricus
(Mollusca:Cephalopoda) in aquaria. Australian Journal of
Marine and Freshwater Research 28:45-56.

Lane, F. W. 1957. Kingdom of the octopus; the life-history of the
Cephalopoda. Jarrolds, London. 287 pp.

Mangold, K. and S. v. Boletzky. 1973. New data on reproductive
biology and growth of Octopus vulgaris. Marine Biology
19:7-12.

Mather, J. A. 1982. Choice and competition: Their effects on occupan-
cy of shell homes by Octopus joubini. Marine Behavior and
Physiology 8:285-293.

Nixon, M. 1969. The lifespan of Octopus vulgaris Lamarck. Pro-
ceedings of the Malacological Society of London 38:529-540.

Perrier, E. and A. T. Rochebrune. 1894. Sur un Octopus nouveau
(O. digueti) de la basse Californie, habitants les coquilles

48 AMER. MALAC. BULL. 6(1) (1988)

des mollusques bivalves. Comptes Rendus de |/Académie
Sciences Paris 118:770-773.

Roper, C. F. E., M. J. Sweeney and C. E. Nauen. 1984. Cephalopods
of the world. Food and Agriculture Organization (FAO) species
catalogue Vol. 3. FAO Fisheries Symposium (125) 3:1-277.

Smale, M. J. and P. R. Buchan. 1981. Biology of Octopus vulgaris
off the East Coast of South Africa. Marine Biology 65:1-12.

Van Heukelem, W. F. 1983. Octopus cyanea. In: Cephalopod Life
Cycles Vol. 1, P. R. Boyle, ed. pp. 267-276. Academic Press,
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and other small mammals in southern lowa. Proceedings of
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Wells, M. J. and J. Wells. 1977. Cephalopoda:Octopoda. /n:
Reproduction of marine invertebrates Vol. 4. Gastropods and
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Malacological Bulletin 4:240 (Abstract).

Date of manuscript acceptance: 21 April 1987

RESEARCH NOTE

THE NEED FOR QUANTITATIVE SAMPLING TO CHARACTERIZE
SIZE DEMOGRAPHY AND DENSITY OF FRESHWATER
MUSSEL COMMUNITIES

ANDREW C. MILLER AND BARRY S. PAYNE
U. S. ARMY ENGINEER
WATERWAYS EXPERIMENT STATION
ENVIRONMENTAL LABORATORY
VICKSBURG, MISSISSIPPI 39180-0631, U. S. A.

ABSTRACT

An accurate estimate of density of all mussels in a community, regardless of size, requires
collecting total substratum samples. As part of a monitoring program cn bivalves in large rivers, 0.25
m? quadrat total substratum samples were collected by divers at two dense beds, one in the upper
Mississippi River at Prairie du Chien, Wisconsin, the other in the lower Ohio River near Olmsted,
Illinois. A linear relationship existed between the cumulative number of species obtained and the
logarithm of the number of quadrats sampled. Using this relationship it was estimated that 40 and
200 samples, respectively, were required to accurately assess species richness at high and low densi-
ty sites in the upper Mississippi River. Because of the contagious nature of these beds, reliable densi-
ty estimates for all unionid species required at least 7 to 12 quantitative samples. Dominant species
were characterized by infrequent but fairly strong recent recruitment, illustrating the necessity of col-
lecting and processing total substratum samples to obtain juveniles.

An evaluation of the condition of a mussel bed should
be based upon measurements of species richness, relative
abundance, density, and recruitment. Accurate determination
of all of these parameters, except perhaps species richness,
requires that quantitative samples of bottom material be ob-
tained and sieved for all live mussels regardless of size.
Although this approach is used in most benthic surveys, it
is rarely applied in studies of mussels in large rivers. In these
habitats mussels often occur in substratum too consolidated
to allow quantitative sampling using devices such as Ponar,
Eckman, Peterson, or Shipek dredges (Isom and Gooch,
1986). The Surber sampler (Henderson, 1949) and suction
pumps (Mattice and Bosworth, 1979) have been used to quan-
titatively collect bivalves in shallow streams. However, the oc-
currence of unionids in deep water and in consolidated gravels
has made quantitative studies of these communities difficult.

Brails, or crowfoot dredges, were developed by com-
mercial fishermen and have been used to study the distribu-
tion and relative abundance of unionids in large rivers (e.g.
Smith, 1898; Baker, 1903; Coker, 1918; Starret, 1971), but
surveys conducted with these devices suffer numerous and
variable biases (e.g. Scruggs, 1960; Krumholz et al., 1970;

Thiel et a/., 1980; Kovalak et a/., 1986). Semi-quantitative
surveys have been performed by having divers equipped with
SCUBA retrieve mussels by feeling for them within quadrats
(e.g. Duncan and Thiel, 1983; Isom and Gooch, 1986; Kovalak
et al., 1986) or along transects (e.g. Brice and Lewis, 1979;
Isom and Gooch, 1986). Search and feel methods are almost
certainly biased against species characterized by small-sized
animals or juveniles of species characterized by large-sized
animals, but have improved our understanding of mussel
distribution in large rivers relative to use of brails (e.g. Isom
and Gooch, 1986; Kovalak et a/., 1986).

The purpose of this paper is to describe a sampling
approach that utilizes quantitative substratum removal to ac-
curately assess size demography and density of unionids in
large river habitats. These studies were conducted as part
of a monitoring program on population and community struc-
ture of bivalves at prominent beds to assess impacts of water
resource development.

STUDY SITES

Studies were conducted at two mussel beds, one

American Malacological Bulletin, Vol. 6(1) (1988):49-54

49

50 AMER. MALAC. BULL. 6(1) (1988)

located in the east channel of the upper Mississippi River near
Prairie du Chien, Wisconsin (RM 636) and the other in the
lower Ohio River near Olmstead, Illinois (RM 967). Both beds
were several km long, at least 300 m wide, and were found
in stable substratum. Sampling sites were in fairly deep water
(4-6 m at typical low water levels in early fall), and located
a minimum of 100 m from the periphery of the bed. Mussel
beds were identified from published information (e.g. Havlik
and Stansbery, 1978, for the site on the upper Mississippi River
and Williams, 1969, for the site on the lower Ohio River). The
approximate size of each bed, and location of sites was deter-
mined by a diver performing a general reconnaissance. A
mussel bed was defined as a contiguous area of stable sub-
stratum where densities were at least 10 individuals per m2.

In the east channel of the Mississippi River in October
1984, 10 samples were taken at each of five sites that were
separated by about 1 km. In July 1985, 30 samples were taken
at each of two sites that consisted of three subsites (see
Hurlbert, 1984) sampled 10 times each. Preliminary sampl-
ing at these and other beds indicated that at least ten samples
would be required to estimate species richness and total
mussel density. Study sites were separated by a distance of
0.5 to 1.5 km; subsites were within 50 m of each other. In ad-
dition, a pair of quadrats were taken every 1.2 m along a 14.4m
transect in a dense part of the bed.

At the mussel bed in the Ohio River, four sites that were
about 50 m apart were sampled six times in September 1983.
In October 1985 a single site was sampled 13 times, and in
September 1986, eight sites were sampled eight times and
one site was sampled four times. The 1983 and 1985 surveys
were conducted in the upstream half of the bed; the 1986
survey was conducted near the downstream limit of the
bed.

LOWER OHIO RIVER
30

f
7
Y = 7.632 + 9.987 LOG X 7 wo
R2 = 0.974 e, 7 7
(98 IND/m2) yy, yo
20 7 7
7
7
7
y
7
7

Y = 2.058 + 9.388 LOG X
e R2 = 0.902
(35 IND/m2)

CUMULATIVE NUMBER OF SPECIES

10 100
CUMULATIVE NUMBER OF 0.25 m2 SAMPLES

METHODS

At each site in a bed, a diver collected samples from
within an aluminum 0.25 m2 quadrat that was positioned in
a haphazard manner near an identifying buoy. The diver
transferred all substratum, which included sand, gravel, shells,
and live organisms, from each quadrat into a 20 / bucket. In
consolidated gravel, digging tools were needed to remove all
material to a depth of 10-15 cm. Collection of a single sample
required 5-15 min. The bucket was pulled or winched to the
surface and transported to shore, where collected material
was washed through a graduated series of sieves. The finest
sieve had a mesh aperture of 4 mm. Material retained on each
screen was examined for live mussels; 5-15 min were required
to wash and pick each sample. Collected mussels were taken
to a mobile laboratory, identified by species, and their shell
lengths measured to the nearest 0.1 mm. Individuals not need-
ed for voucher specimens were returned to the river.

Although the dive crew consisted of 3 - 4 individuals,
only a single diver worked a site at a time. Support person-
nel consisted of 4 - 6 individuals that helped position boats
and transport and process samples. Depending upon logistics
and experience of personnel, 10 - 30 samples were collected
and processed to completion each day.

RESULTS

SPECIES RICHNESS AND RELATIVE ABUNDANCE.

The cumulative number of species obtained at any site
was a linear function of the logarithm of the number of
quadrats sampled. This relationship is portrayed for represen-
tative high and low density sites located in beds in the Ohio
and Mississippi rivers (Fig. 1). The dashed lines in figure 1

UPPER MISSISSIPPI RIVER

7
VG /
y, 7
7 7

Y =9.666+12599LOGX / ~

R2 = 0.965 re 7

(149 IND/m2) a

7
7
7
7
7
7

Y = 1.895 + 12.342 LOG X
R2 = 0.964
(27 IND/m2)

10 100

Fig. 1. Cumulative number of species in relation to number of quadrat samples collected in the Ohio and upper Mississippi rivers.

MILLER AND PAYNE: QUANTITATIVE SAMPLING FOR MUSSELS oil

extend the number of samples beyond that which was col-
lected in these surveys to a value necessary to obtain all
species reported to exist in the beds in the Ohio (Miller et a/.,
1986) and Mississippi rivers (Havlik and Stansbery, 1978).
Extensions of these lines are not statistically valid in a strict
sense. However, such extensions are instructive and sup-
ported by the ubiquity of relationships between estimates of
species richness and the number of samples collected
(McNaughton and Wolf, 1973). These relationships depict the
diminishing rate of addition of new species as more samples
are taken. For example, at a high density site in the Ohio River,
15, 21, 24, and 25 species were yielded by 10, 20, 40, and
60 samples, respectively. At a dense site in the Mississippi
River, all species known from this reach of the river were col-
lected with 40 samples; however, approximately 200 samples
would be needed at the low density site to obtain all species
present (Fig. 1).

A large number of quadrats must be sampled to ob-
tain all species in both beds because most mussels are locally
uncommon. Both beds were heavily dominated by a single
unionid species. For example, Amblema plicata (Say) com-
prised 54.3% of the east channel community in the upper
Mississippi River in 1985 and Fusconaia ebena (Lea)
represented 66.7% of all native unionids in the Ohio River
in 1985. Of the 29 species collected in the upper Mississippi
River in 1985, 16 accounted for less than 1% of the community.
Lampsilis higginsi (Lea), a species on the Federal list of en-
dangered species, ranked 17th on the list and comprised 0.61
and 0.58% of the community in 1984 and 1985, respectively.
Of the 23 species collected in the Ohio River during the 1985
survey, 11 accounted for less than 1% of all native unionids.
Plethobasus cooperianus (Lea), a federally-listed endangered
species was collected in this bed using qualitative techniques
(Miller et a/., 1986), but was not obtained in quadrat samples
during any year.

DENSITY

In the upper Mississippi River, the average density of
all unionid species at the five sites sampled in 1984 and the
two sites (consisting of three subsites) sampled in 1985 ranged
from 22 + 20 to 202 + 36 individuals per m2 (+ standard
deviation, N = 10 at each site or subsite). In the lower Ohio
River, densities ranged from 47 + 24 to 80 + 20(N = 6)
in 1983, and 102 + 30(N = 13) in 1985, and 9 + 3 to 31
+ 6 individuals per m2 (N = 8 for eight sites and N = 4 for
one site) in 1986.

A further illustration of the contagious nature of these
molluscan communities is shown by the results of sampling
along a transect within the bed in the Mississippi River (Fig.
2). The spatial heterogeneity of the bed directly affects the
number of samples required to accurately estimate mussel
density with a defined level of accuracy and precision. The
number of samples required to estimate mussel density was
determined by treating a set of replicate samples within a site
as a pilot survey. To determine the number of quadrats
necessary to achieve a desired precision of total mussel den-
sity a procedure from Green (1979:41) was used. This requires
making an estimate of the mean and standard deviation of

sles MUSSELS
20 -
N
—€
isk
3
2
SPECIES
10
5 =
0 [oe | ee, S| ae eet | | ey ees)
0 1 2 3 4 5 6 7 8 9 10 11 12 «#13
Site No. 1.5m

Fig. 2. Total individuals and species richness (pool for two 0.25 m2
quadrats) from a transect in the upper Mississippi River.

the population from preliminary sampling. The number of
samples necessary to achieve a desired estimate of preci-
sion is a function of the variance of the pilot sample. For each
of the 11 site-specific surveys in the upper Mississippi River
we computed the number of samples necessary to estimate
the average density of all mussels within either 10 or 30%
of the actual average density with a 5% probability of be-
ing incorrect. We found that from 1.4 to 37.5 (mean = 12.2)
samples were required to be within 30% and from 12.9 to 246.5
(mean = 109.8) samples were required to estimate to within
10% of the actual average density of all unionids at the 11
sites. The coefficient of variation of density estimates was
lower at sites in the Ohio River than those in the upper
Mississippi River. From 1.7 to 15.9 (mean = 6.2) samples were
required to be within 30%, and from 19.2 to 143.0 (mean =
55.9) samples were needed to estimate to within 10% of the
average density for the 14 sites in the lower Ohio River.

SIZE DEMOGRAPHY

The most useful aspect of quantitative sampling was
the detection of patterns in population recruitment for
Amblema plicata in the Mississippi River and Fusconaia ebena
in the Ohio River. The dominant species in both beds showed
evidence of tremendous annual variation in recruitment
strength. Mature females of both species produce glochidia
each year for many years during their reproductive life span,
and survival of glochidia through metamorphosis and settle-
ment is contingent upon a number of abiotic and biotic
variables. Thus, large annual variations in recruitment should
be expected in such populations.

Shell length frequency histograms for Amblema plicata
in the Mississippi River indicate that recruitment success was

52 AMER. MALAC. BULL. 6(1) (1988)

Fusconaia ebena

Amblema plicata

LOWER OHIO RIVER, 1983 UPPER MISSISSIPPI RIVER, 1985

(N = 78)
100

80
E
E
x 60
-
1)
2
uw
: U
4
— 40
uu
x
n

Ge camer

O
0 20%

(N = 85) (N = 190) (N = 194)

0
CA

OO

i

10%

PERCENT FREQUENCY

Fig. 3. Representative length-frequency histograms for Fusconaia ebena at two sites in the Ohio River in 1983 and Amblema plicata at two

sites in the Mississippi River in 1984.

low for year classes represented by mussels between 40 and
60 mm in 1984 (Fig. 3). It is unlikely that selective mortality
of these size classes occurred in the post-settlement stage
of the life cycle. Also, recruitment exhibited spatial variability
within the mussel bed. Although the sites in the Mississippi
River depicted in figure 3 were less than 1 km apart, recruit-
ment rates were not uniform throughout the mussel bed.

Intersite differences in patterns of size demography
were not detected for Fusconaia ebena in the Ohio River (Fig.
3). However, evidence of annual variation in recruitment was
more striking for F ebena in the Ohio River (Fig. 3). In this
population a single year class (probably 1982), represented
by mussels 16-20 mm long, accounted for 70% of all in-
dividuals of this species collected in 1983. This same year
class remained a dominant feature of the size demography
of this population when assessed again in 1985 and 1986 (Fig.
4, cohort centered at 29 mm in 1985 and at 36 mm in 1986).
Strong recruitment was not observed for any year class since
1982.

DISCUSSION

The areas studied in the upper Mississippi River near
Prairie du Chien, Wisconsin and the lower Ohio River near
Olmsted, Illinois are among the most dense and rich mussel
beds in these two rivers (Havlik and Stansbery, 1978; Miller
et al., 1986). Rigorous quantitative sampling at both beds
revealed common features of community and population
structure. Both communities are marked by heavy dominance

LOWER OHIO RIVER

100 1983 1985 1986
(n = 256) (n = 252) (n = 93)

| :

SHELL LENGTH, mm
8 8
in]
N
oO
o
— -

PERCENT FREQUENCY

Fig. 4. Annual variation in recruitment for Fusconaia ebena in the
lower Ohio River in 1983, 1985, and 1986.

by a single species and a large number of uncommon species.
This same pattern is observed in most natural communities
(e.g. Hughes, 1986). Based upon results of these studies,

MILLER AND PAYNE: QUANTITATIVE SAMPLING FOR MUSSELS 53

mussels in large rivers are no exception to this general rule.

Quantitative samples are required for unbiased
estimates of the relative abundance of species. A conse-
quence of the local rarity of many unionids is that a large
number of quantitative samples are required to obtain all
species at a site (see also Kovalak et a/., 1986). A combina-
tion of qualitative and quantitative sampling methods is the
most efficient way to completely assess community composi-
tion. Qualitative surveys facilitate estimation of species
richness, and quantitative surveys are required for estimation
of relative species abundance.

Density of mussels is estimated with fewer samples
than species richness and relative abundance. Based on our
results, seven to twelve quadrat samples were sufficient to
estimate the average density within 30% of the actual average
density at a site with a 5% probability of being incorrect.
As these statistics demonstrate, intersite variation in average
density and the coefficient of variation of density estimates
can be substantial. This is a direct consequence of the con-
tagious nature of these communities and illustrates the need
for a study design which includes adequate number of sites
and replicates. Intrasite variation could be reduced by collec-
ting individual samples within cells of a large (4m x 4m)
16-celled PVC grid secured to the bottom with pins. This pro-
cedure could help to eliminate diver bias and could reduce
the coefficient of variation of estimates made of particularly
contagious distributions.

Annual and intersite variation in recruitment was evi-
dent in both mussel beds. Intersite variation in patterns of size
demography, like intersite variation in density, argues for
sampling replicate sites. Annual variation in recruitment of
dominant mussels, while evident in both beds, was particularly
striking for Fusconaia ebena in the lower Ohio River. The size
demography of this species was such that a single year class
will remain a dominant feature of the size structure of this
population for years hence.

Most riverine unionids have a long life span, take
several years to mature, and appear to have great annual
variation in recruitment success. These organisms are
especially sensitive to commercial fishing and development
of water resource projects. Regulation of commercial harvests
and protection of habitat must be based on knowledge of
population and community demographics. Currently we are
conducting annual surveys at important mussel beds to
monitor long-term trends in these parameters. However, most
mussel studies in large rivers have not been sufficiently quan-
titative to elucidate important aspects of the biology of these
invertebrates. Judgments on the condition of freshwater
bivalves in large rivers should be based on quantitative
substratum sampling that enables accurate determination of
relative abundance, density, recruitment, growth, and
mortality.

ACKNOWLEDGMENTS

The tests described and resulting data presented herein,
unless otherwise noted, were obtained from research conducted
under the Environmental Impact Research Program and the En-

vironmental and Operational Water Quality Studies Program, of the
United States Army Corps of Engineers by the U. S. Army Engineer
Waterways Experiment Station. In addition, funds were provided by
the U. S. Army Engineer Districts Louisville and St. Paul. The follow-
ing divers participated in this work: Larry Neill, Roger W. Fuller, William
H. Host, Jr., and Jim Walden, Tennessee Valley Authority; Ron Fet-
ting, Robert Sikkla, Ed Stran, and Bill Wolf, U. S. Army Engineer
District, St. Paul. Assistance in the field was provided by C. Rex
Bingham, Waterways Experiment Station, Vicksburg, Mississippi;
Kevin Cummings, Illinois Natural History Survey, Champaign, Illinois;
Paul Hartfield, Mississippi Museum of Natural Science, Jackson,
Mississippi; Robert King, Central Michigan University, Mt. Pleasant,
Michigan; Teresa Naimo, Tennessee Technological University,
Cookeville, Tennessee; Robert Read, Wisconsin Department of
Natural Resources, Madison, Wisconsin; Terry Siemsen, U. S. Army
Engineer District, Louisville, Kentucky; Robert Simmonet, Minnesota
Museum of Natural Science, St. Paul, Minnesota; Carl Way, North-
western University, Chicago, Illinois; Dan Wilcox, U. S. Army Engineer
District, St. Paul, Minnesota. Permission was granted by the Chief
of Engineers to publish this information.

LITERATURE CITED

Baker, F. C. 1903. Shell collecting in Mississippi. Nautilus 16:102-105.

Brice, J. and R. Lewis. 1979. Mapping of mussels (Unionidae) com-
munities in large streams. American Midland Naturalist
101:454-455.

Coker, R. E. 1918. Freshwater mussels and mussel industries of the
United States. Bulletin of the Bureau of Fisheries 36:11-89.

Duncan, R. E. and P. A. Thiel. 1983. A Survey of the Mussel Den-
sities in Pool 10 of the Upper Mississippi River. Technical
Bulletin No. 139 of the Wisconsin Department of Natural
Resources, Madison, Wisconsin. 14 pp.

Green, R. H. 1979. Sampling Design and Statistical Methods for En-
vironmental Biologists. John Wiley and Sons. New York. 257 pp.

Havlik, M. E. and D. H. Stansbery. 1978. The naiad mollusks of the
Mississippi River in the vicinity of Prairie du Chien, Wiscon-
sin. Bulletin of the American Malacological Union for 1977:9-12.

Henderson, C. 1949. Value of the bottom sampler in demonstrating
the events of pollution on fish-food organisms in the Shenan-
doah River. Progressive Fish Culturist 11:217-230.

Hughes, R. G. 1986. Theories and models of species abundance.
American Naturalist 128:879-899.

Hurlbert, S. H. 1984. Pseudoreplication and the design of ecological
field experiments. Ecological Monographs 54:187-211.

Isom, B. G. and C. Gooch. 1986. Rationale and sampling design for
freshwater mussels, Unionidae, in streams, large rivers, im-
poundments, and lakes. In: Rationale for Sampling and Inter-
pretation of Ecological Data in the Assessment of Freshwater
Ecosystems. B. G. Isom, ed. pp. 46-59. American Society for
Testing and Materials, STP 894, Philadelphia, Pennsylvania.

Kovalak, W. P., S. D. Dennis and J. M. Bates. 1986. Sampling effort
required to find rare species of freshwater mussels. /n: Ra-
tionale for Sampling and Interpretation of Ecological Data in the
Assessment of Freshwater Ecosystems, B. G. Isom, ed. pp.
46-59. American Society for Testing and Materials, STP 894,
Philadelphia, Pennsylvania.

Krumholz, L. A., R. L. Bingham and E. R. Meyer. 1970. A survey of
the commercially valuable mussels of the Wabash and White
Rivers of Indiana. Proceedings of the Indiana Academy of
Sciences 79:205-226.

Mattice, J. and W. Bosworth. 1979. Modified venturi suction sampler
for collecting asiatic clams. Progressive Fish Culturist 41:121-123.

54 AMER. MALAC. BULL. 6(1) (1988)

McNaughton, S. J. and L. L. Wolf. 1973. General Ecology. Holt,
Rinehart and Winston, Inc., New York. 710 pp.

Miller, A. C., B. S. Payne and T. S. Siemsen. 1986. Plethobasis
cooperianus. Nautilus 100:14-17.

Scruggs, G. D. 1960. Status of the Freshwater Mussel Stocks in the
Tennessee River. Special Scientific Report - Fisheries No. 370.
U. S. Fish and Wildlife Service, Washington, D.C. 41 pp.

Smith, H. M. 1898. The mussel fishery and pearl-button industry of
the Mississippi River. Bulletin of the U. S. Fish Commission
43:289-314.

Starret, W. C. 1971. A survey of the mussels of the Illinois River, a
polluted stream. Illinois Natural History Survey Bulletin

30:265-403.

Thiel, P., M. Talbot and J. Holzer. 1980. Survey of mussels in the up-
per Mississippi River Pools 3 through 8. /n: Proceedings of the
UMRCC Symposium on Upper Mississippi River Bivalve
Mollusks, Upper Mississippi River Consortium Conservation
Committee. J. L. Rasmussen, ed. pp. 148-156. Rock Island,
Illinois.

Williams, J. C. 1969. Project Completion Report for Investigation Pro-
jects: Mussel Fishery Investigations, Tennessee, Ohio and Green
Rivers. Kentucky Department of Fish and Wildlife Resources,
Lexington, Kentucky. 107 pp.

Date of manuscript acceptance: 7 May 1987

SYMPOSIUM ON THE BIOLOGY
OF THE POLYPLACOPHORA

ORGANIZED BY
ROBERT C. BULLOCK
UNIVERSITY OF RHODE ISLAND

AMERICAN MALACOLOGICAL UNION

KEY WEST, FLORIDA
21 JULY 1987

55

ANCESTORS AND DESCENDENTS: RELATIONSHIPS
OF THE APLACOPHORA AND POLYPLACOPHORA

AMELIE H. SCHELTEMA

BIOLOGY DEPARTMENT
WOODS HOLE OCEANOGRAPHIC INSTITUTION
WOODS HOLE, MASSACHUSETTS 02543, U. S. A.

ABSTRACT

Four organ systems, pericardium of primitive mollusks, shell ontogeny and spicule formation
in chitons and aplacophorans, chaetoderm oral shield, and aplacophoran radula, are described and
their relationships discussed. The discussion suggests: (1) a coelomate ancestor of the mollusks; (2)
a polyphyletic origin of shell, one for Conchifera and another for chitons; (3) a single class Aplacophora
containing two taxa, the Chaetodermomorpha and Neomeniomorpha; (4) an archimolluscan radula
with a pair of separate radular membranes bearing rows of single teeth. Evidence is presented that
contradicts the following hypotheses: (1) an acoelomate origin of mollusks; (2) the division of
aplacophorans into two classes; (3) the derivation of the univalved molluscan shell from a common
stem with the eight-shelled chitons. The concept of a subphylum Aculifera is rejected as unnecessary

since it holds no essential information.

Hypotheses of early molluscan evolution in the last fif-
teen years have proposed an acoelomate, turbellariomorph
pre-molluscan ancestor with a mucoid dorsal cover and a
broad, ciliated locomotory sole through which opened a mouth
(Fig. 1) (Salvini-Plawen, 1972, 1980, 1985; Haas, 1981; Boss,
1982; Poulicek and Kreusch, 1983; see also Fretter and
Graham, 1962; Stasek, 1972). According to such theories, this
pre-mollusk gave rise to an archimollusk with a spiculose in-
tegument, an unpaired radular membrane, and a mouth that
opened through the ventral locomotory surface. The archi-
mollusk then gave rise to two major taxa, the burrowing
aplacophorans (Chaetodermomorpha = Caudofoveata) and
an ‘‘adenopod’’, with seven transverse rows of scales and a
head separated from the sole. The second group of aplaco-
phorans, the footed Neomeniomorpha (= Solenogastres
sensu Salvini-Piawen), have split off from the hypothetical
“‘adenopod’’, the latter giving rise to an ‘‘archiplacophoran”’
with plates formed from coalesced scales. The ‘‘archiplaco-
phoran’”’ in turn was the precursor of the Polyplacophora on
one hand and the rest of the shelled mollusks, the Conchifera,
on the other (for recent accounts and bibliographic references,
see Runnegar and Pojeta, 1985; Wingstrand, 1985; Salvini-
Plawen, 1985). The subphylum Aculifera, recognized by Haas
(1981) and formerly, but no longer, by Salvini-Plawen (cf. 1972,
1980), includes the extant Aplacophora and Polyplacophora
as well as the hypothetical archimollusk, adenopod and arch-
iplacophora; all other mollusks form the subphylum Con-
chifera. Salvini-Plawen (1980) considers the Chaetoder-

CONCHIFERA
Archiconchifera aS =
Se
a
eae ACULIFERA

4
/ Polyplacophora

-

Neomeniomorpha
(Solenogastres sensu
Salvini-Plawen)

_ as

Chaetodermomorpha
(Caudofoveata)

(1) Archiplacophora

(2) Adenopod

(3) Archimollusk

~ =

Se i

(4) Turbellariomorph

Fig. 1. Phylogeny of the Mollusca (adapted in part from Salvini-
Plawen, 1980; Haas, 1981; Poulicek and Kreusch, 1983). Questioned
in the text is the validity of: (1) an archiplacophoran origin of the
Conchifera; (2) separation of the aplacophoran taxa Chaetodermo-
morpha and Neomeniomorpha by the existence of an Adenopog; (3)
an archimolluscan radula with an undivided radular membrane; (4)
an acoelomate ancestor. Compare with figure 14.

American Malacological Bulletin, Vol. 6(1) (1988):57-68

oy,

58 AMER. MALAC. BULL. 6(1) (1988)

momorpha to belong to the subphylum Scutopoda; all remain-
ing mollusks, including the Neomeniomorpha, constitute the
subphylum Adenopoda.

Evidence presented here draws on recent observations
or experiments on shell and radula formation, the structure
of the oral shield of the burrowing aplacophorans, and the
size of pericardial spaces in three primitive molluscan classes.
The evidence raises questions about the validity of four
hypotheses: (1) there is a monophyletic (archiplacophoran)
origin of chitons and conchiferan mollusks; (2) the two
aplacophoran taxa belong to two separate classes; (3) the
most primitive molluscan radula had an undivided radular
membrane; (4) the ancestor of mollusks was acoelomate

(Fig. 1).

SHELL AND SPICULES

APLACOPHORA AND POLYPLACOPHORA

The Aplacophora and Polyplacophora have been
classified together either as the Amphineura because of their
similar ladder-like nervous systems (not examined here), or
as the Aculifera because of their similar integumental struc-
tures: papillae, spines, and cuticle. Indeed, these anatomical
relationships between the two groups have been used to
justify the inclusion of Aplacophora within the Mollusca (for
historical reviews, see Hyman, 1967; Scheltema, 1978),

although they are better regarded as symplesiomorphic traits,
shared primitive states that do not necessarily show close
evolutionary relationships.

Beedham and Trueman (1968) found similarities in the
histochemistry of aplacophoran and chiton integumental cuti-
cle and concluded that ‘‘the cuticle of the Aplacophora is ten-
tatively equated with an early mucoid stage in the evolution
of the molluscan shell... [The cuticle of Acanthochiton] has
in addition a discrete inner cuticular layer which may act as
a semi-conducting membrane in the deposition of calcareous
plates’ (p. 443). The papillae of Aplacophora and Poly-
placophora are probably homologous (F. P. Fischer, pers.
comm.); the papillae and aesthetes of Polyplacophora are
likewise homologous (Fischer et a/., 1980; Fischer, 1988).

The process of calcareous spicule formation, most
recently investigated by Haas (1981), is alike in aplacophorans
and chitons (Fig. 2). In both taxa, a spine is secreted extra-
cellularly within an invagination of a single cell. A basal cell
secretes calcium carbonate, and as the spicule grows beyond
this cell, a crystallization chamber is sealed off by a collar
of neighboring cells. The megaspines in chitons, which do
not occur in Aplacophora, are formed by a proliferation of the
original single basal cell.

The attempt to find further similarities in calcium car-
bonate deposition that would link the Aplacophora and
Polyplacophora by examining embryogenesis has led to less
conclusive comparisons. Larval development in the two

Fig. 2. Spicule formation in Aplacophora and Polyplacophora. A. Primitive Neomeniomorpha. B. Lepidochitona cinerea (Linnaeus). An organic
pellicle has not been demonstrated around spicules of the Aplacophora. (After Haas, 1981.) (b, basal cell; n, neighboring cell; p, organic pelli-

cle; s, spicule). Scale bars = 1 mm.

SCHELTEMA: APLACOPHORA AND POLYPLACOPHORA 59

Fig. 3. Reported ontogeny in an aplacophoran, Nematomenia
banyulensis Pruvot, and a chiton, Lepidochitona corrugata Reeve [=
Middendorffia caprearum (Scacchi)]. A. Pruvot’s larva, a single obser-
vation, lateral view, of a metamorphosing larva of Nematomenia with
seven dorsal calcareous ‘plaques’, slightly imbricated and formed
of rectangular, plainly juxtaposed spicules”’ (translated from Pruvot,
1890). The larva did not survive to a juvenile stage. B. Defective shell
formation in Lepidochitona corrugata (= Chiton polii (Philippi) as il-
lustrated by Kowalevsky (1883) with separate granules of calcium car-
bonate deposited along seven plate fields. Coalescence of these
granules does not lead to normal growth of shell plates (see Kniprath,
1980). C. Birefringence under cross-polarized light in a normally
developing Lepidochitona corrugata larva. Noncalcareous areas are
stippled; the birefringent spicular girdle and six straight, uninterrupted
anlagen of the shell plates are without stippling, as are the birefringent
rosette-shaped larval eyes. (A and B after Salvini-Plawen, 1972: Fig.
29, after comparison with the original drawings of Pruvot, 1890, and
Kowalevsky, 1883; C drawn after photograph by Kniprath, 1980: Fig.
1b.). Scales not known.

groups is dissimilar, but Salvini-Plawen [1972, 1980, 1985 (with
qualifications)] argues for homology between seven rows of
spicules seen once in a single aplacophoran larva
[Nematomenia banyulensis Pruvot, Pruvot (1890)] and the
development of shell in the larva of the chiton Lepidochitona
corrugata (Reeve) (= Chiton polii Philippi) by a coalescence
of granules (Fig. 3A, B) (Kowalevsky, 1883). The rows of
spicules observed by Pruvot have not subsequently been seen
in any other aplacophoran larvae [Epimenia verrucosa
(Nierstrasz), Halomenia gravida Heath, Neomenia carinata
Tullberg; see Hadfield (1979) for a summary]. Pruvot’s draw-
ing is a lateral view, and the often-copied dorsal view show-
ing seven rows of spicules is a hypothetical reconstruction
(Salvini-Plawen, 1972; Wingstrand, 1985).

Recently, Kniprath (1980) reported from rearing ex-
periments that in the larvae of both Lepidochitona corrugata
[=Middendorffia caprearum (Scacchi] and /schnochiton rissoi
(Payraudeau) the anlagen of the plates are secreted as
uninterrupted rods along narrow transverse depressions, the
shell or plate fields, after the development of girdle spicules
(Fig. 3C). When Lepidochitona larvae were reared at
temperatures of 149-16°C, shell development was normai, but
all larvae raised at higher temperatures of 189-21°C were ab-
normal and developed granules similar to those reported by
Kowalevsky (1883). These granules, even when they coa-

lesced, produced defective shell plates.

The seven “‘plaques’’ of Pruvot’s larval aplacophoran
specimen are said to reflect the number of plates in the early
fossil chiton Septemchiton (Hyman, 1967; Salvini-Plawen,
1980) and the seven ‘‘larval’’ plaques of chitons (Salvini-
Plawen, 1985). However, Rolfe (1981) has shown that the most
anterior plate of Septemchiton, a burrowing form, although
greatly reduced is indeed present and that Septemchiton
therefore has a full complement of eight plates. Although the
caudal plate in chitons is usually added last during develop-
ment, sometimes only after an extended period of five weeks
(Pearse, 1979), it is not clear whether this time lapse reflects
an ancestral chiton with only seven plates or is simply a result
of development as a chiton elongates. In many adult aplaco-
phorans with single overlapping layers of flat, leaf-like spicules,
the bases of the spicules are aligned in rows that are
transverse to the long axis of the animal (unpub. data); it
would therefore not be surprising to find spicules lined-up
in metamorphosing larvae that could be mistaken for
‘plaques’.

Evidence for the coalescence of spines is said to be
shown by three sets of broad spicules, or shields, on the head
of the juvenile aplacophoran Nematomenia protecta (Thiele,
1913). This conclusion is based on spicule shape only, without
reference to the underlying epithelium; the number of cells
involved in secreting a ‘‘shield’’, a single cell or more than
one cell, is not known, despite the inferred epithelial connec-
tion constructed by Salvini-Plawen (1985: Fig. 36D). The
evidence for coalescence therefore remains unsubstantiated.

Both aplacophorans and chitons retain in common a
phylogenetically early mode of calcium carbonate deposition
in the form of spicules, but until further observations on
aplacophoran embryogenesis prove to the contrary, close
evolutionary relationship between the formation of
aplacophoran spicules and chiton shells is considered un-
demonstrated. There is no evidence within chitons themselves
that spicules have coalesced to form shell plates.

POLYPLACOPHORA AND THE OTHER SHELLED
MOLLUSKS (CONCHIFERA)

The process of shell formation in chitons is argued here
to be unique among mollusks. In those gastropods, bivalves,
and cephalopods for which the entire shell ontogeny has been
studied, earliest calcium carbonate deposition is preceded,
first, by formation of a shell-field and shell-field invagination
from part of the dorsal ectoderm and, second, by the secre-
tion of an organic pellicle, usually equated with periostracum,
over the invagination (Fig. 4A) (Kniprath, 1981; Eyster and
Morse, 1984). [In the Cephalopoda, yolk interferes with in-
vagination and, instead, ectoderm builds up in an elevated
ring (Kniprath, 1981)]. Calcium carbonate is then secreted
beneath the organic pellicle. In the nudibranch Aeolidia
papillosa (Linnaeus), the early organic pellicle is overlain by
long cytoplasmic processes that presumably seal off the
crystallization chamber under the pellicle (Fig. 4B) (Eyster and
Morse, 1984).

In chitons, no shell field invagination forms (Fig. 4C).

60 AMER. MALAC. BULL. 6(1) (1988)

Deposition of a shell plate anlage takes place within a
transverse depression bounded and sealed off by long,
overlapping microvilli that lie beneath a gelatinous mucoid
substance, certainly not periostracum, and questionably
equated with a cuticle (Fig. 4C, D) (Kniprath, 1980; Haas et
al., 1980; Haas, 1981).

Not only are the ontogenetic processes of shell forma-
tion different in chitons and the Conchifera, but structures of
the fully formed shells are also unlike and homologies are
difficult to discover. Periostracum in the Conchifera, a struc-
ture conservative in manner of its secretion and in composi-

Fig. 4. Larvel shell deposition in (A, B) the gastropod Aolidia papillosa
(Linnaeus) and (C, D) the chiton /schnochiton rissoi (Payaudreau).
In A, an organic pellicle (arrows) covers the lumen of the shell field
invagination (L); in B, the edge of the pellicle can be seen to be
overlain by a cytoplasmic extension (e). Calcium carbonate has not
yet been deposited. (Drawn after photographs in Eyster and Morse,
1984: Figs. 1, 2). In C, calcium carbonate of the shell plate (p) has
been deposited under the overlapped microvilli (s, ‘‘stragulum’’); a
mucus layer (m) covers the stragulum. In D, microvillar processes
(s) have pulled apart and a cuticle (c) with a contrasted outer layer
is beginning to form; M is perhaps a mucus cell (C and D after
Kniprath, 1980.) Scale bars: A = 10 um; B = 0.5 um; C; D approx-
imately 6 pm.

tion (Grégoire, 1972), does not exist in chitons, although Haas
(1981) has demonstrated the presence of a thin cuticle, or pro-
periostracum, overlying the tegmentum and a properiostracal
groove surrounding each shell plate. There is no nacreous
layer in chiton shells as found in other mollusks, and the cross-
lamellar structure of the shell plates is crystallographically uni-
que, with bundles of crystal fibers in the lamellae ordered so
that their c-axis ‘‘coincides with the bisectrix of these cross-
ing fibers’’ (Haas, 1981: 403) and the ‘‘whole complex acts
crystallographically as a single crystal’ (Haas, 1977: 392). In
other molluscan cross-lamellar structures, the angle between
crystal fibers is about 110°; in gastropods they lie between
90°-130° (Wilbur and Saleuddin, 1983). Haas (1981) considered
the cross-lamellar structure of chitons to be homologous with
the nacreous layer of other shelled mollusks and imagined
that both arose from an undifferentiated inner layer of the
“‘archiplacophoran’”’ plates. The shell of the Conchifera
became univalved he believed by fusion of the shell and shell
fields. There is no evidence, however, that the dynamics in-
volved in the process of earliest shell deposition through the
interplay of shell-field invagination and pellicle in Conchifera
could have evolved from the very different process of shell-
plate production found in chitons.

Thus, recent work on the ontogeny and structure of
shell in chitons and Conchifera shows such major differences
between them that it can be questioned whether there was
a monophyletic origin of molluscan shell, or rather one origin
for chitons and a second for the remaining extant and extinct
Conchifera. Tubules in the shells of the monoplacophoran
Neopilina (Schmidt, 1959), bivalves (e.g. Waller, 1980), and
gastropods have sometimes been considered homologous
with the aesthete canals of chitons and argued as a support
for a monophyletic origin of molluscan shell (e.g. Salvini-
Plawen, 1985), but the homology is so far uncertain. When
the ontogenetic development of Neopilina becomes known,
perhaps a basis will be found for deciding whether molluscan
shell has a monophyletic or polyphyletic origin.

CHAETODERM ORAL SHIELD AND
THE ARCHIMOLLUSK

One of the original arguments for dividing the
Aplacophora into two classes and, ultimately, into two sub-
phyla depends on the hypothesis that mollusks have a
turbellariomorph, or flatworm, ancestry. This phylogeny is
based on a supposed homology and similarity in mode of
locomotion between mollusks and flatworms by means of a
“ventral mucociliary gliding surface’ (Salvini-Plawen, 1972,
1980: Fig. 5, 1985; see also Trueman, 1976). The molluscan
archetype, like the flatworms, is said not to possess a Separa-
tion of the head from the foot, and the mouth consequently
opens through the sole; innervation of the sole is said to be
from both the cerebral ganglia and ventral nerve cord. [Stasek
(1972) has illustrated but not discussed a head separate from
the locomotory sole in the turbellariomorph molluscan
precursor.]

Support for the flatworm-like archimolluscan locomo-

SCHELTEMA: APLACOPHORA AND POLYPLACOPHORA 61

tory ventral surface is said to be shown by the cerebrally in-
nervated oral shield of the burrowing Chaetodermomorpha
(= Caudofoveata) (Fig. 6A); that is, the shield is regarded as
a remnant of the original gliding surface (Salvini-Plawen,
1972, 1980, 1985). The homology with a creeping sole was
originally based on histologic similarities in the morphology
and arrangement of nerve and mucous cells that lie in the
epidermis beneath the oral shield cuticle of chaetoderms and
the spiculeless cuticle within the foot-furrow of the creeping
neomeniomorphs [Hoffman, 1949; for a translation and ex-
planation, see Scheltema (1983)]. The homology, however, is
spurious since molluscan ectoderm, with or without cuticle,
is richly supplied with both nerve and mucous cells. Further-
more, Salvini-Plawen (1985) has described (but not illustrated)
the specialized ultrastructure of the oral shield, consisting of
interdigitated microvilli with glycocalyxes and supporting
fibers.

The oral-shield cuticle and epithelium in six genera
(Scutopus, Limifossor, Prochaetoderma, Metachaetoderma,
Falcidens, and Chaetoderma) representing all families of
chaetoderms are continuous with pharyngeal (oral tube) cuti-
cle and epithelium (Scheltema, 1981, 1983). Light microscopy
does not reveal a border where the oral shield cuticle joins
the pharyngeal cuticle (Figs. 5, 6B), but ultrastructural studies
would define this area better. Scutopus is considered to be
the most primitive chaetoderm because of its least differen-
tiated midgut (Scheltema, 1981) and because of the evidence
of ventral fusion of the cuticle (Salvini-Plawen, 1972). In this
genus only scattered pyriform mucous cells open through the

Fig. 5. Oral shield of a Chaetodermomorpha: section through the
mouth, pharynx, and oral shield of Scutopus megaradulatus Salvini-
Plawen showing continuous cuticle of pharynx and oral shield (from
650 m off Cape Hatteras, North Carolina, U. S. A., 34°14.8’N,
75°46.7’W; fixed in formalin, preserved in alcohol, stained with
haemotoxylin/Gray’s double contrast, sectioned at 0.7 yum.) (c,
spiculose cuticle of integument; n, nerve fibers from precerebral
ganglion; 0, cuticle of oral shield; p, cuticle of pharynx). Small arrow
indicates change from oral shield cuticle with a thickened outermost
layer to homogeneous cuticle of pharynx. Scale bar = 0.05 mm.

Fig. 6. Oral shield of Scutopus megaradulatus. A. Anterior view of
oral shield in situ surrounding darkened mouth in center. B.
Semischematic drawing of area between large arrowheads in figure
5 showing histology of pharyngeal and oral shield cuticle (lettering
and small arrow as in Fig. 5). Scale bars: A = 0.3mm; B = 0.05mm.

oral shield, further refuting Hoffman’s homology, which likened
the lobes of mucous cells opening at the lateral edges of the
oral shield in advanced Chaetodermatidae with the pedal
gland of Neomeniomorpha. This important aspect of Hoff-
man’s homology linking lobed mucous cells of the oral shield
and foot furrow was ignored by Salvini-Plawen (1980) while
retaining the homology itself. Definitive evidence that the oral
shield is a part of a vestigial ventral sole would require inner-
vation from the ventral (= pedal) nerve cord rather than from
the cerebral ganglia.

Thus, the oral shield of the Chaetodermomorpha is
considered here to be an autapomorphy, a cerebrally inner-
vated external continuation of pharyngeal cuticle like a lip
belonging to the head, not to a ventral sole. There is no con-
vincing evidence that it is a remnant of an original creeping
sole homologous to the ventral surface of a turbellarian flat-
worm. The separation of the Aplacophora into two classes
based on the supposed (1) plesiomorphy of ventral innerva-
tion of the chaetoderm oral shield by the cerebral ganglia and
(2) apomorphy of a head separate from the foot in the
neomenioids and all other mollusks except chaetoderms is
unsatisfactory. A head separate from the foot is considered
here to be a plesiomorphy shared by mollusks generally but
lost in the bivalves and, because of their burrowing habit, also
in the chaetoderms.

62 AMER. MALAC. BULL. 6(1) (1988)

RADULA

APLACOPHORAN RADULA

Evidence from the radula morphology of aplacophor-
ans and from the ontogeny of gastropod and chiton radulae
suggests that the molluscan radula orginated as a paired
structure.

The radula in chitons, the monoplacophoran Neopilina,
gastropods, and scaphopods is a chitinous structure formed
of a single continuous ribbon, or radular membrane, which
bears serial rows of teeth; both ribbon and teeth are continual-
ly secreted at the proximal end of a pharyngeal diverticulum,
the radular sac (Fretter and Graham, 1962; Kerth, 1983;
Scheltema, unpub. data). Each row of teeth has left and right
sides and usually a central, or median, tooth. The radula is
bilaterally symmetrical around the central tooth, that is, the

teeth of each side are mirror images of one another. Along
the length of the ribbon each tooth has the same shape as
the tooth in front of and behind it, that is, the rows of teeth
are serially repeated.

In the Aplacophora, the radula is formed in the usual
manner and is likewise bilaterally symmetrical and serially
repeated (Figs. 7A, 8A, C). The radula has been called
monostichous or monoserial if there is only a single tooth in
a row; with two mirror-image teeth in each row, distichous or
biserial; and with more than two mirror-image teeth,
polystichous or polyserial (Nierstrasz, 1905).

The usual type of radula in the Aplacophora is
distichous; a central tooth is lacking in nearly all species. Uni-
que among mollusks the radular membrane itself is divided
down the middle so that the entire radula is a bipartite,
bilaterally symmetrical, serial structure consisting of two strips

Fig. 7. Aplacophoran radula of Simrothiella species. A. Simrothiella sp. b (undescribed); at left are the newest, proximal teeth and fused radular
membrane (arrow); distally (on the right) the membrane is bipartite and spirals ventrally down into two ventral pharyngeal pockets. B. Close-up
of fused, proximal end of radula shown in A. (Whole amount in glycerine; see Scheltema, 1981, for dissecting technique). C. Simrothiella sp.
a (undescribed), sagittal section through one side of radula, indicated by single arrowheads; double arrowheads show radula within the ventral
pharyngeal pocket (Specimens from 2,633 m at 20°50’N, 109° 0.6’W; sections treated as in Fig. 5). Scale bars: A = 100 um; B = 30 um;

C = 100 um.

SCHELTEMA: APLACOPHORA AND POLYPLACOPHORA 63

of continuous ribbon, each strip with rows of single denticulate
teeth which are the mirror image of the opposed teeth (Figs.
7A, 8A, C). The two parts of the radular membrane are fused
to a greater or lesser extent lengthwise along their medial (in-
ner) edges forming a one-piece, unipartite radular ribbon
along part of its length (Figs. 7B, 8A; Scheltema, 1981).
The structure of the radula is clear only when it is
dissected and isolated from surrounding tissue (Scheltema,
1981). Reconstructions from histologic sections have resulted

y
BY DB

By)

ote

WY
Sy

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=D)
It

5
K

TE
SY 5
HA

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i]

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r)

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FZ
rel

Fig. 8. Radula of Simrothiella sp. b (undescribed), radular membrane
indicated by stippling. A. Entire radula of a juvenile specimen, dor-
sal view, anterior (oldest teeth) at top. Teeth of only left half of radula
shown; teeth on the right are the mirror-image of those on the left.
Denticles are added to the teeth medially as the radula widens and
lengthens. B. Distal, oldest part of left radular strip shown folded under
in A from ventral pharyngeal pocket; original, first-formed tooth is
retained. C. Two views of the same two adjacent teeth from an adult
specimen: upper teeth drawn in dorsal view as if they were on the
right side of the radula, medial denticles on left; lower teeth from left
side of radula drawn from beneath radular membrane. D. Most anterior
part of the same adult radula from which teeth in C were drawn; com-
parison with juvenile radula B indicates that there is dissolution at
the distal end of the radula within the ventral pharyngeal pocket
(Specimens from 2,633 m at 20°50’N, 109°06’W). Scale bars in mm.

in misconceptions of actual structure and probable modifica-
tions during its evolution [e.g. Nierstrasz, 1905; Salvini-Plawen,
1972, 1978 (Simrothiella), 1985].

In order to differentiate the two states that exist for the
radular membrane among mollusks, the terms “‘bipartite’” and
“‘unipartite’’ are used here, and the terms using ‘—stichous”’
are reserved for descriptions of the radular teeth only. Thus,
a distichous radula can be either uni- or bipartite, but a
monostichous radula is necessarily unipartite. The terms with
‘serial,’ which should mean ‘‘arranged in series,’ are not
used here, thus obviating the confusion of such a descrip-
tion as ‘“‘monoserial with paired teeth.’

As in other radulate Mo!lusca, the radular membrane
in Aplacophora appears to migrate forward as teeth are add-
ed by the odontoblasts; in most species the membranes turn
anteroventrally into paired or unpaired ventral pharyngeal
pockets, where dissolution of the radula apparently occurs
(Figs. 7C, 8D). Unlike grazing gastropods and chitons, in all
but one family of Aplacophora the teeth show no wear and
thus do not rasp.

The entire radula of juvenile specimen of Simrothiella
(0.9 mm in length) has been examined. Within each ventral
pharyngeal pocket is preserved the earliest ontogenetic
development; the first tooth is a nondenticulate bar on a wide
expanse of radular membrane (Fig. 8B). As the radula grows
in length and width, denticles are added to the teeth medial-
ly, i.e. at their inner edges (Fig. 8A). Histologic cross-sections
through the proximal, blind end of the radular sac show odon-
toblasts in two discrete groups, each presumably bound by
basement membrane (Figs. 9, 10). The two groups lie within
a single sac, surrounded in the usual manner by muscle.

Within the Aplacophora, the radula has evolved at least
twice from having a bipartite, distichous radula (Figs. 7, 8) to
a radula with a unipartite radular membrane. In the Donder-
siidae (Fig. 11), the radula is altogether absent or consists of

Fig. 9. Radular sac of Simrothiella sp. a (undescribed). Anterior view
of somewhat oblique cross-section through proximal end showing
membranes (arrowheads) bounding right and left groups of radula
secretory cells (Specimen from 2,633 m at 20°50’N, 109°06’W). Scale
bar = 35 um.

64 AMER. MALAC. BULL. 6(1) (1988)

Fig. 10. Semischematic representation of radular sac cross-section
shown in figure 9 (er, epithelium of radular membrane; m, membranes
bounding left and right groups of radula secretory cells; mu, mus-
cle; od, odontoblasts; t;, early tooth, or perhaps denticle, not yet stain-
ing with haemotoxylin; tz, older tooth stained by haemotoxylin). Scale
bar = 35 um.

only a few rows of single teeth, usually 6 or fewer. Its
monostichous form appears to be the result of reduction and
fusion of a distichous radula, with two of its paired denticles
fused at tip and base. In the Prochaetodermatidae, the radula
has evolved into a rasping structure with a unipartite radular
membrane and a central tooth, or plate (Fig. 12) (Scheltema,
1981,1985).

There are no distinctive radula characteristics, syn-
apomorphies, held in common or uniquely by the Aplacophora
and Polyplacophora, the latter with rows of usually 17 teeth
on a unipartite radular membrane.

ONTOGENY OF GASTROPOD AND CHITON
RADULAE

Vestiges of an original distichous molluscan radula ex-
ist in the ontogenetic development of the chiton, pulmonate,
opisthobranch, and prosobranch radula. The details of the
developing chiton radula are treated by Eernisse and Kerth
(1987) and Kerth (this symposium). The radula starts as rarely
one to uSually three pairs of lateral teeth on a unipartite radular
membrane with a central tooth added later. In the ontogenetic
development in five families and seven species of pulmonates,
the radula begins as a distichous structure with two
longitudinal rows of lateral teeth on a unipartite radular mem-
brane; further laterals are then added, and finally a central
tooth, which originally may be paired, is secreted thereby
uniting the cross-rows (Kerth, 1979). Pruvot-Fol (1926) figured
the earliest radular teeth of the opisthobranch Polycera,

Fig. 11. Monostichous aplacophoran radula of an undescribed species
of Atlantic Dondersiidae, four aspects; radular membrane not shown.
One denticle is missing from the teeth in the lower two drawings
(Specimen from 805 m, 39°51.3’N, 70°54.3’W). Scale in mm.

Fig. 12. Undivided, unipartite radular membrane of an undescribed
species of Prochaetodermatidae; view of ventral surface (Specimen
from 1,624 m 10°30.0’N, 17951.5’W). Scale = 250 um.

SCHELTEMA: APLACOPHORA AND POLYPLACOPHORA 65

distichous with a ‘‘gouttiere’’ between them. The radular sac
in the opisthobranch Rhodope (Riedl, 1960) and in the
pulmonate Physa (Wierzejski, 1905) originates as a pair of in-
vaginations. In Rhodope, lacking a radula, the paired invagina-
tions are lost; in Physa, they unite to form a single sac. The
developing radular sac in prosobranchs is often bifid (Fretter
and Graham, 1962: 173).

To summarize, the most generalized aplacophoran
radula is unique because it has a bipartite radular membrane
with distichous teeth. Distichous teeth on a unipartite radular
membrane exist ontogenetically in other molluscan groups.

PERICARDIUM

The pericardium is a space lined by mesoderm aris-
ing embryologically from cell 4d; therefore, it may be con-
sidered to be coelom. Raven (1966) questioned, however,
whether coelomic cavities among mollusks arise from
mesodermal bands (schizocoels) as they do among the an-
nelids. [For an extensive overview of gonopericardial com-
plexes within mollusks, see Wingstrand (1985)].

Salvini-Plawen (1968) hypothesized that the pericardial
space evolved within the mesenchyme after the heart, sur-
rounding it and thereby improving its function. Stasek (1972:
Fig. 1A, B) illustrated such a situation in the molluscan precur-
sors. Although the pericardium is relatively small in most
gastropods and bivalves, in the three primitive classes
Aplacophora, Monoplacophora, and Polyplacophora it is
spacious relative to the size of the heart (Fig. 13). In Neopilina
the pericardium is paired, and in the aplacophoran Chaeto-
dermomorpha and most Neomeniomorpha it has either small
or large, paired lateral extensions (‘‘horns’’ in early literature),
whose function is not known. Ontogenetically, in the single
species of aplacophoran for which size during development
is mentioned (Baba, 1938), the pericardium is already large
before the heart develops.

How the pericardium functionally could have evolved
in a pre-mollusk as a small space, then have become spacious
and probably paired, and finally again become reduced in
size, is difficult to imagine. Moreover, during organogenesis,
the pericardium develops before the heart and the heart arises

Fig. 13. Heart and pericardium in the primitive molluscan classes Aplacophora (A, B), Monoplacophora (C), and Polyplacophora (D) showing
large pericardial spaces in relation to the size of the heart. In B, C, and D the heart is stippled and the pericardium is blank. A. Chaetoderma
nitidulum Loven, sagittal section through pericardium, heart, and gonopericardial duct (after Scheltema, 1972). Paired auricles (a) open into
the ventricle on each side of an atrioventricular valve (avwv). Gonads empty through paired ducts (g) into the pericardium (pc), and coelomoducts
(cd) lead from the pericardium to the cloaca (not shown). The large paired lateral extensions of the pericardium (e) are known as ‘‘horns’’
in the older literature. B. Simrothiella sp. a (original drawing), same specimen as in figure 9. Somewhat oblique cross-section through the pericardium
(pc), ventricle (v), and lateral extension of the pericardium (e). C. Neopilina galatheae Lemche, dorsal view (after Lemche and Wingstrand,
1959). The pericardium (pc) and ventricles (v) are paired; two pairs of auricles (a) open into each ventricle. It is not known whether there is
a connection between the pericardia and gonads (see Wingstrand, 1985). D. Acanthopleura echinata, dorsal view (after Plate, 1898). Two pairs
of ostia (0) open on each side into the ventricle (v); the number of ostia varies from one to four pairs, according to species (a, auricle; ab,
aortal bulb; aw, atrioventricular valve; cd, coelomoduct; e, lateral extension of pericardium; g, gonopericardial duct; 0, opening between auri-
cle and ventricle; pc, pericardium; v, ventricle). Scales not indicated.

66 AMER. MALAC. BULL. 6(1) (1988)

from the dorsal or inner epithelium of the pericardium (Baba,
1938; Raven, 1966), suggesting that evolution of the pericar-
dium probably preceded that of the heart. The large pericar-
dial spaces in the Aplacophora, Monoplacophora, and Poly-
placophora point to a coelomate rather than to an acoelomate,
turbellariomorph ancestor and lead one to re-examine the
evidence for ancestral relationship between the annelids and
mollusks (see Vagvolgyi, 1967; Wingstrand, 1985).

DISCUSSION

ACOELOMATE VERSUS COELOMATE
MOLLUSCAN ORIGINS

The hypothesis that the ancestor of mollusks was
acoelomate is rejected in favor of a coelomate origin because:
(1) primitive molluscan taxa have large pericardial spaces; (2)
evidence is lacking that the pericardial space began as a small
opening in mesenchyme lined by mesoderm; (3) Wingstrand’s
evidence (1985) strongly suggests a molluscan ‘‘derivation
from advanced oligomeric Spiralia (‘proto-annelids’ or ‘proto-
articulates’)’’ (p. 8) (Fig. 14).

The existence of large pericardial spaces in the
primitive extant mollusks has not been considered in
hypotheses of an acoelomate molluscan origin. Rejection of
the hypothesis of reduced metamery as the origin of
molluscan coelom is probably correct (Salvini-Plawen, 1968);
however, one need not suppose, therefore, a total absence
of either coelom or metamery. Reiger (1985), after careful com-
parative studies of the fine structure of acoel connective tissue,
argued that the acoelomate Bilateria themselves are derived
through progenesis from a coelomate ancestor.

SHELL AND SPICULES

The Aplacophora probably evolved from a shell-less
rather than from a shelled ancestor. Evidence for this asser-
tion comes from properties of the cuticle (see SHELL AND
SPICULES above) and from a comparison of numbers of dor-
soventral muscles that run between the outer body wall and
foot among various mollusks. In the Neomeniomorpha, two
bilateral sets of oblique bands are repeated serially along the
body; they are considered homologous to the dorsoventral
pedal muscles in other mollusks (Salvini-Plawen, 1972). The
evolution of dorsoventral musculature, which coevolved with
the shell, has been toward reduction in number, from eight
in Polyplacophora and tryblidian Monoplacophora to one in
most Gastropoda. The serial arrangement of numerous bands
in the Neomeniomorpha is considered therefore to be a
plesiomorphy that preceded shell development and its con-
sequent reduction of dorsoventral musculature.

No convincing published evidence links the process
of extracellular spicule formation by a single cell (Haas, 1981)
with the development of shell fields and shell deposition. The
only common attribute of spicule and shell formation is that
both are extracellular deposits of calcium carbonate.

Three types of calcium carbonate coverings are found
in the Mollusca: spicules in Aplacophora and Polyplacophora;
the shell plates of the Polyplacophora with a thin

POLYPLACOPHORA CONCHIFERA

Polyplacophora

Conchiferan
ancestor

ye

Testacean ancestor

APLACOPHORA (no shell)
Chaetodermomorpha Neomeniomorpha
(Caudofoveata) (Solenogastres)
Common aplacophoran Adenopod?

ancestor

Coelomate (oligomerous?)
molluscan ancestor

Fig. 14. Phylogeny of the Mollusca (adapted from Wingstrand, 1985).
The questioned Adenopod can be dropped (see argument in sec-
tion ‘‘Chaetoderm oral shield and the archimollusk’’). The text raises
questions about a common testacean ancestor in comparing chiton
and conchiferan shell formation and structure (see argument in sec-
tion ‘Shell and Spicules’’). A coelomate molluscan ancestor, whether
or not oligomerous, is corroborated here (see section ‘‘Pericardium’’).
A common aplacophoran ancestor descended directly from the stem
mollusk is indicated (See sections ‘‘Chaetoderm oral shield and the
archimollusk’’ and ‘‘Aplacophora, a monophyletic group’). The stem
mollusk had a paired radula with a two-part radular membrane and
distichous teeth (see section ‘‘Radula’’).

(nonperiostracal) organic cover, tegmentum, and
hypostracum; and the conchiferan shell with periostracum,
prismatic layer, and nacreous layer. The trend has been to
treat these calcium carbonate structures as homologous, with
a morphocline leading from spicules to plates by coalescence
in chitons (e.g. Salvini-Plawen, 1972), and from the 8 shell
fields in chitons to the single shell field of univalves and
bivalves (e.g. Haas, 1981). From the evidence of structure and
ontogeny, and discounting the problematic ‘‘Pruvot’s larva,’
the existence of this morphocline is seriously questioned.

Is there a single ancestor for polyplacophorans and
the remaining shelled mollusks? Wingstrand (1985) makes a
strong case for such a hypothetical testacean ancestor,
equivalent to the archiplacophoran of figure 1, based on
synapomorphies of radula with its supports and musculature,
oral flaps, digestive system, pharyngeal diverticula, 8 pairs
of pedal retractors, and, possibly, the number and position

SCHELTEMA: APLACOPHORA AND POLYPLACOPHORA 67

of atria (Fig. 13). The shells in chitons are considered to be
autapomorphies, but the shell fields and the mineralization
process are homologous and monophyletic in chitons and
Conchifera. Reasons have been stated above (section on
Shell and Spicules) for doubting this homology (Fig. 14).
Answers to questions about Pruvot’s larva and the relation-
ship of polyplacophoran plates to conchiferan shells could
lie in the unknown embryology of Neopilina and with the yet-
to-be reexamined Pruvot’s larva.

RADULA

The direction of evolutionary change in the structure
of the aplacophoran radula appears to be from a paired, or
bipartite, radular membrane to a single, unipartite ribbon. The
rationale for this polarity is based on several points. (1) Rasp-
ing seems a more advanced, complicated function for a radula
over a simple ability to grasp as found in most Aplacophora.
Rasping probably requires the integration of structure provid-
ed by a unipartite radular membrane. Only among the Pro-
chaetodermatidae is there wear of the anterior teeth, i.e.
evidence of rasping (Scheltema, 1981, 1985), and here the
radular membrane is also unipartite. (2) All other radulate
aplacophorans except the Dondersiidae and Chaetodermatide
with reduced and specialized teeth (Fig. 11; Scheltema, 1972)
have a bipartite radular membrane with a fused, unipartite
section that often retains visible evidence of fusion; the region
of this fused section is not fixed but varies among families
and genera (Scheltema, 1981). It is possible, but not parsimon-
ious, to imagine that the radular membrane was originally
unipartite, then divided into two, and finally fused again;
however, if so, the odontoblasts producing such a secondari-
ly derived, paired radula would have to evolve from a single
into a paired group of cells. (3) During ontogeny of the radula
in chitons and gastropods, the central tooth is added only after
several rows of one or more pairs of lateral teeth have been
formed. Presumably the median part of the ribbon is where
an originally paired ribbon became unified; subsequently
odontoblasts for the central tooth could come into being.

The paired structure of the aplacophoran radula is con-
sidered to be the primitive form in mollusks because the direc-
tion of evolution, distichous bipartite to distichous unipartite
in Aplacophora, is continued in the ontogeny of the gastropod
radula, from distichous unipartite to polystichous. Since
aplacophorans probably evolved from a shell-less ancestor
(see above), the distinctive molluscan structure of a radula
was already present when shell evolved (Fig. 14). The
aplacophoran plesiomorphic bipartite radula does not form
a basis for linking the Aplacophora closely to any other tax-
on of mollusks.

APLACOPHORA, A MONOPHYLETIC GROUP

The Aplacophora should not be separated into two
classes or subphyla on the erroneous homology of the
chaetoderm oral shield with a turbellariomorph creeping sole.
The. oral shield is an autapomorphy of the Chaetodermo-
morpha. The Neomeniomorpha and Chaetodermomorpha
form a monophyletic group with the following probable syna-
pomorphies: a rounded worm shape; a dorsoterminal sen-

sory organ [a chemoreceptor lying external to the mantle cavi-
ty, and not known to be ontogenetically or functionally
homologous to the osphradium within the mantle cavity of
other mollusks (Haszprunar, 1987)]; three to six pairs of
precerebral ganglia or swellings (Salvini-Plawen, 1978, 1985);
a reproductive system in which the gonads empty into the
pericardium through gonopericardial ducts and the pericar-
dium is emptied into the cloaca through coelomoducts (Fig.
13A) (but see Salvini-Plawen, 1972, 1985). An adenopod
ancestor becomes a superfluous construct (Fig. 14). As the
direction of evolution of organ systems within the Aplacophora
becomes clear, new insights into the evolution of mollusks
should come to light.

ACKNOWLEDGMENTS

The opportunity to bring together some thoughts on the im-
portance of Aplacophora in the phylogeny of Polyplacophora and the
Mollusca generally was provided by the kind invitation of Robert C.
Bullock to participate in the chiton symposium of the American
Malacological Union, July 1987. Douglas J. Eernisse, Janice Voltzow,
Klaus Kerth, and Rudolf S. Scheltema receive my heartfelt thanks
for reviewing the manuscript, which underwent some vigorous
rewriting following their suggestions. | am grateful to them for rescu-
ing me from circular reasoning, imprecise terminology, redundan-
cies and unfinished thoughts, but the errors which remain are mine
alone.

Figures 7 through 12 are based on specimens collected under
National Science Foundation grant numbers GA-31105 and
OCE-8025201.

This paper is contribution number 6599 of the Woods Hole
Oceanographic Institution.

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Date of manuscript acceptance: 19 October 1987

THE GILLS OF CHITONS (POLYPLACOPHORA)
AND THEIR SIGNIFICANCE IN MOLLUSCAN PHYLOGENY

W. D. RUSSELL-HUNTER
DEPARTMENT OF BIOLOGY, SYRACUSE UNIVERSITY,
SYRACUSE, NEW YORK 13244, U. S. A.

and

MARINE BIOLOGICAL LABORATORY
WOODS HOLE, MASSACHUSETTS 02543, U. S. A.

ABSTRACT

It was demonstrated in 1965 that gills of chitons are not paired structures but are added during

growth and can show asymmetry. More recent studies, largely on living Chaetopleura apiculata (Say)
at Woods Hole, confirm the broad homologies of each chiton gill with the aspidobranch ctenidium re-
tained in several stocks of Archaeogastropoda. In particular, similar organization is found of afferent
and efferent blood vessels in the gill axis; of alternating ctenidial leaflets; and of lateral, frontal, and
abfrontal cilia. In addition to like ciliary functions, both the gastropod aspidobranch gill and each in-
dividual chiton gill show similar neuromuscular reflexes in cleansing mucus-bound sediment. One dif-
ference, due to the functional organization of each row of chiton gills into a pallial curtain dividing the
mantle groove, is the occurrence of Velcro-like ciliary junctions. Unlike junctions in mytilid and other
““filibranch’’ bivalves, which are modified lateral cilia linking adjacent filaments on the same gill, these
ciliary junctions link leaflets on adjacent gills and probably represent modified frontal cilia. The coor-
dinated and dynamic functioning of this ctenidial curtain is emphasized, and it is suggested that the
adaptive basis on which chitons evolved a curtain by replicating gills, rather than by elongation of ctenidial
parts, results from the dynamic pallial groove (unlike the fixed shapes of pallial cavities in bivalves
and shelled gastropods). Otherwise chiton gills, along with those of protobranchiate bivalves and cer-
tain archaeogastropods, are little altered from ‘‘archetypic’’ molluscan ctenidia.

All archetypes are speculative, available as temporary models of ancestors to be tested by predic-
tions and retrodictions. However, data on gills and other replicated structures in chitons (like data on
Neopilina, and on molluscan capacity for degrowth) appear to exclude hypotheses involving true

metameric segmentation from models of ancestral molluscs.

The multiplied organ systems found in chitons have
to be considered in any discussion of metamerism in primitive
molluscs. It was demonstrated several years ago (Russell
Hunter and Brown, 1965) that the gills of chitons are not paired
structures but are added singly during growth, with the result
that several species show asymmetry in ctenidial numbers
between the left and right sides of individual chitons. Gills
continued to be added in adults to meet increased respiratory
needs with growth of live tissue mass, and it was concluded
that the rows of ctenidia, and probably the other multiplied
structures in chitons, reflect functional replication (Russell
Hunter and Brown, 1965) rather than the vestiges of more ex-
tensive ancestral segmentation as assumed by Lemche
(1959b, 1966). The significant feature of the gill rows in dividing
the mantle grooves of chitons into functionally inhalant and
exhalant chambers had been elucidated by Yonge (1939), and

this also stressed functional rather than vestigial multiplica-
tion of the gills. Since the discovery in 1952 of the living mono-
placophoran genus, Neopilina (Lemche, 1957; Lemche and
Wingstrand, 1959), discussion of possible metamerism in
primitive molluscs has been revised, and continues into the
1980’s. Recent Russian investigators of the multiplied struc-
tures of chitons (Minichev and Sirenko, 1984) have again con-
cluded that there is no evidence of annelid-like metamerism
in their morphogenesis. In his most recent, and beautifully
detailed, account of anatomy in Monoplacophora, Wingstrand
(1985) still concludes that in chitons, ‘‘an oligomeric repeti-
tion, probably 7- or 8-metamerism is present”’ (p. 87, see also
pp. 77-81). Given the currency of such divergent views, it
seemed appropriate to use this symposium on the Biology
of Polyplacophora to present some more recent observations
on the functioning of the gills in living chitons. These studies

American Malacological Bulletin, Vol. 6(1) (1988):69-78

69

70 AMER. MALAC. BULL. 6(1) (1988)

were mostly carried out with Chaetopleura apiculata (Say) at
Woods Hole.

The material presented here involves not only the func-
tional morphology of individual ctenidia in living chitons, but
also their combined dynamics as a gill curtain. Two general
aspects will be emphasized. First, each chiton gill is a true
ctenidium, structurally and functionally homologous with the
aspidobranch gill in certain archaeogastropods and with the
more primitive gills of protobranchiate bivalves. In addition
to reviewing the integrated ciliary and circulatory functions,
new observations are presented on neuromuscular cleans-
ing reflexes common to all these primitive molluscan ctenidia.
Secondly, new observations give emphasis to the coordinated
functioning of the replicated gills as a ctenidial curtain dividing
the inhalant from the exhalant pallial chambers, but con-
forming dynamically to the changing shape and hydraulics
of each pallial groove. Some speculation on this as the likely
adaptive basis for gill replication in chitons follows, along with
a discussion of these and other multiplied structures of
chitons. Finally, the implications of such functional replica-
tion are considered in relation to hypotheses on interrelation-
ships among the major classes of molluscs, and on metameric
segmentation in models of ancestral molluscs.

MATERIALS AND METHODS

In 1979-80 and again in 1986-87, living specimens of
Chaetopleura apiculata (Say) were studied at the Marine
Biological Laboratory (MBL), Woods Hole. This is the ‘‘Com-
mon Eastern Chiton’’ of the Atlantic seaboard of the north-
eastern United States, and most of the material came from
boulders on the Buzzards Bay side of Penzance Point near
Woods Hole. Other early observations on gills in living
specimens of Lepidochitona cinerea (L.) were carried out in
1961-63 in Scotland. Over the years 1961-87, other casual
observations on living chitons have been made on Tonicella
marmorea (Fabricius) and Acanthochitona crinita (Pennant)
in Scotland, and on T. rubra (L.) in Massachusetts and Maine.
The only observations on Lepidopleurus cancellatus (Sower-
by) and L. asellus (Gmelin) were on material already fixed.

Most observations were made under dissecting micro-
scope (at magnifications from X7 to X30) using incident
lighting, with living chitons crawling inverted under glass
slides, or on the convex sides of watch glasses. A few obser-
vations utilized a temporary ‘‘inverted microscope’ arrange-
ment of a dissecting microscope pod to check on water cur-
rents in chitons crawling dorsal side up (that is with pallial
grooves and their contained ctenidia directed downwards).
Elucidation of water and ciliary currents, and mapping of
mucous secretion and accumulation, involved the injection
of particles into the pallial grooves. Particles used included
fine carborundum, carmine, Ankolor scarlet S, and dried milk
powder. The three figures are diagrams, admittedly reduc-
tionist cartoons, each derived from sets of many sketches.
Figure 1 is basically from Lepidochitona, and figures 2 and
3 from Chaetopleura. Some specimens were preserved after
partial narcotization using propylene phenoxetol (for details
of this method, see Russell Hunter and Brown, 1965), fixa-

tion in 12% formalin in sea water, and storage in 10% glycerol.
Temporary microscope mounts were made of individual gills,
both living and fixed, for viewing both by incident and by
transmitted light.

OBSERVATIONS

GENERAL ARRANGEMENT AND
NUMERICAL ASYMMETRY

In chitons, the mantle cavity is in the form of two nar-
row pallial grooves running between the foot and the broad
mantle edge or girdle on each side. Each pallial groove con-
tains a row of gills, the bases of which are attached deep in
the groove on the girdle side (Fig. 1). The gill curtain forms
a functional division of the pallial groove longitudinally into
an inhalant chamber, ventral on the girdle side, and an ex-
halant chamber placed dorsally and pedally (Fig. 1B). As in
all molluscan mantle complexes, the anus along with kidney
and genital openings discharge in the exhalant stream. Newly
formed ctenidia are at the anterior end of each row (Fig. 1A).
As growth continues in adult chitons, ctenidia are added
anteriorly, irregularly and independently on each side.
However, for any species of chiton, there is always a broad
correlation between gill number and adult tissue mass
(Russell Hunter and Brown, 1965). Asymmetry in ctenidial
numbers between the left and right sides of individual chitons
occurs in most chiton species. For populations of four chiton
species studied in detail, the percentages of asymmetric in-
dividuals were 19.5%, 46.3%, 48.4% and 69% (Russell Hunter
and Brown, 1965). In most species the numbers of individual
chitons with extra left gills are apparently balanced by the
numbers with extra right gills. However, Gowlett-Holmes and
Zeidler (1987) have described a new species, Acanthochitona
saundersi, for which all available specimens have 11 ctenidia
on the right side and 10 ctenidia on the left side. Asymmetries
of ctenidial numbers have been found in at least fifteen
species of chitons, and could well occur in the majority of
chiton species (Minichev and Sirenko, 1984; A. M. Jones, pers.
comm.).

CTENIDIAL FUNCTIONAL MORPHOLOGY

When the gills of a living chiton are viewed from the
ventral side, the free tips are seen to be directed toward the
edge of the foot (the inner wall of the pallial groove). The gills
bulge convexly toward the observer and their axes are de-
fined by the prominent efferent branchial vessels (Fig. 2, ebv).
The other (exhalant) face of each axis contains the narrower
afferent branchial vessel. The leaflets, which alternate on
either side of the gill axis, are short and wide (almost semicir-
cular in face view, Fig. 3), and their tips are opposed (one
to one, or one to two) to the tips of leaflets on the next
ctenidium in the row (Fig. 2).

Water is moved dorsally (and pedally) by broad bands
of lateral cilia (which are more flagella-like) toward the inner
and posteriorly directed exhalant chamber (Fig. 3), in a
physiologically efficient counter-flow to the blood circulation
(afferent branchial vessel to efferent branchial vessel) within

RUSSELL-HUNTER: GILLS OF CHITONS 71

Fig. 1. Diagrams of the mantle groove in a chiton (based on Lepidochitona) showing (A) ventral aspect of anterior part of groove, and (B) cross-
section of the groove at a central ctenidium. Note that inhalant part of the groove (INH) is girdle-ventral and exhalant part (EXH) is pedal-dorsal

(gi, girdle; ft, foot; pg, pallial groove).

each ctenidial leaflet. On the inhalant (INH) side the edges
of the leaflets bear shorter frontal cilia (that is, on the facing
edges of Fig. 2), and on the exhalant (EXH) side the leaflet
edges bear abfrontal cilia. Both frontal and abfrontal cilia have
a cleansing (particle-moving) function rather than water pro-
pulsion, and transport particles around the leaflet edges
toward the axis. The free tips of each leaflet bear specialized
longer, less motile cilia that entangle in a Velcro-like fasten-
ing (x on Fig. 3) with the corresponding cilia on the leaflet
tips of the adjacent ctenidium. From their position, and
development in ctenidial buds, these ciliary junctions linking
adjacent gills probably represent modified frontal cilia.
The assemblage of microstructures and their functions
shown by the chiton gill are thus essentially similar to those
found in the primitive ‘‘aspidobranch’”’ plume gill of the Ar-
chaeogastropoda. If an individual chiton gill is specifically
compared with the single plume gill in the limpet, Acmaea
testudinalis (Muller), the only significant difference involves
the Velcro-like ciliary junctions on the chiton leaflet tips. There
are obviously minor differences of microanatomy such as the
outline proportions of the leaflets, and the distribution of lateral
cilia on the leaflet faces, but these seem trivial in comparison
with the broader concert of structures and functions. The gill
axes with alternating leaflets are essentially identical in ar-
rangement, as are the dorsal afferent branchial vessel and
the ventral efferent vessel carrying oxgenated blood back to
the heart. The lateral, frontal and abfrontal cilia are arranged
in the same way and, in both, the lateral cilia produce a flow
of water through the gill (and through the mantle cavity) in
the opposite direction to the blood flow. Chiton gills are true
Ctenidia, structurally and functionally homologous with those
of other molluscs. The rows of chiton gills are clearly not
neomorphic structures, secondary respiratory organs as in

some marine limpets like Patella, or in various groups of
freshwater pulmonate snails (Russell-Hunter, 1978; McMahon,
1983), but have to be regarded as rows of multiplied ctenidia.

CTENIDIAL CLEANSING REFLEX

Surprisingly little attention has been paid to the
muscular movements of primitive molluscan ctenidia. A
relatively new set of observations on chiton gills concerns the
fact that each ctenidium can move in a patterned cleansing
reflex. To anticipate a little, the sequence of movements in
the individual chiton ctenidium seems to be exactly similar
to that in the cleansing ‘‘flick”’ of the single plume gill in forms
like Acmaea.

In the axis of the chiton ctenidium, longitudinal mus-
cle fibers lie around and below the two major blood vessels.
When both sets of muscle strands contract together, the gill
is shortened and pulled toward its base, with a consequent
decrease in the gill’s contained blood volume. Gill retraction
of this sort can be accomplished in 0.2 to 0.8 seconds. Re-
extension of the gill is always slower (Several seconds) with
blood being passed in hydraulically by action of distant an-
tagonists. If the muscle under the afferent branchial vessel
alone contracts (stretching the muscle on the efferent side)
then the gill curls up into the pallial groove, the ctenidial tip
moving away from the foot (Figs. 1, 2). In the opposite case,
if the muscle under the efferent branchial vessel contracts
the whole gill is straightened and its tip could hit the foot edge
or the substratum-surface or both.

If the cleansing cilia (frontal) are experimentally load-
ed by introducing material (suitably dense but small, like fine
grade carborundum) onto the inhalant face of the gill, the
foreign particles become mucous-bound and are moved

pe AMER. MALAC. BULL. 6(1) (1988)

ft
gl

Fig. 2. Diagrammatic view of four ctenidia in Chaetopleura from the
ventral inhalant side (INH). The axes show the efferent branchial
vessels (ebv), and there are frontal cilia on the facing edges of the
leaflets. Note that the tips of leaflets are opposed one to one, or one
to two (INH, inhalant current; gi, girdle; ft, foot; pg, pallial groove).

toward the axial ciliary tract (Fig. 3) and thence toward the
gill tip. In a healthy chiton, accumulation of this sort at the
tip provokes a reflex action sequence. The reflex is not gravi-
ty dependent and can be observed in chitons in all postural
relations to the horizontal. The same reflex takes place if
foreign material is loaded on the abfrontal (exhalant) face of
the gill. The patterned cleansing reflex occurs in three sequen-
tial phases. First, for two to three seconds, more blood is
pushed in while the gill expands. (It is difficult to measure this,
but the overall volume increase at this phase is usually be-
tween 20% and 50%). Secondly, the muscle strands under
the afferent branchial vessel contract relatively slowly, taking
between 2 and 5 seconds. Thirdly, the muscle under the ef-
ferent branchial vessel contracts relatively rapidly, taking be-
tween 0.1 and 0.2 seconds, and flicks the tip toward the foot
and substratum-surface while simultaneously shortening the
gill. (Only at this stage is the contained blood volume reduced
again.) In most cases a mucous-bound pellet of natural sedi-
ment, or foreign particles, leaves the gill surface and remains
on the cilia of the pedal edge or on the substratum. It should
be noted that there is never any question of ciliary junctions
being formed (even temporarily) between the gill tip cilia and
the pedal cilia.

Despite the subtle differences in ctenidial proportions
noted above, this reflex action of the individual chiton gill (ac-
ting, it seems, in a neuromuscular sense as a peripheral
reflex, or almost as an independent effector system) involves
a patterned sequence exactly following that observed in the

aspidobranch gill of Acmaea. Parenthetically, it is worth noting
one somewhat special case observed in living chitons, con-
cerning the last large gill in Chaetopleura. On several occa-
sions it has been observed to ‘‘flick’’ material right out of the
pallial groove, with its tip passing under the girdle in a tem-
porary (and asymmetric) lifting more like the usual local arch-
ing of the girdle for typical temporary inhalant openings. Of
course, this can only occur because of the distinctly different
siting of that last large gill, with its tip directed posteriorly and
girdlewards instead of toward the midline and foot. In this
respect as others, conditions in the lepidopleurid chitons must
be quite different, but we lack observations of living gill
movements. In typically near-holobranch chitons like
Chaetopleura and Lepidochitona, groups of three or more
Ctenidia can flick together. This leads to the second group
of new observations.

THE COORDINATED CTENIDIAL CURTAIN

Even the casual observer of the underside of a living
chiton can see (Fig. 2) the functional organization of each row
of chiton gills into a pallial curtain dividing the mantle groove
along most of its length into inhalant and exhalant chambers.
This is functionally dependent upon the occurrence of Velcro-
like ciliary fastenings on the leaflet tips of chiton gills. Unlike
the ciliary junctions in mytilid and other ‘‘filibranch’’ bivalves
which are modified lateral cilia linking adjacent filaments on
the same ctenidium, these ciliary junctions in chitons link
leaflets on adjacent gills and probably represent modified fron-
tal cilia. If the filibranch gills typical of mussels, scallops or
oysters are disturbed mechanically, the ctenidial filaments
become tangled and the coordinated filtering and sorting func-
tions are temporarily lost. Given otherwise healthy conditions
and a little time (usually only a few minutes), the filaments
will ‘“‘crawl’’ by ciliary action over each other until the ap-
propriate ciliary junctions are reconnected and the seeming-
ly continuous corrugated lamella re-established as a porous
water-propelling and filtering surface. Similar processes oc-
cur if the ctenidial curtain is mechanically disturbed in a
healthy chiton. Individual gills can carry out slower flicks
across the pallial groove, but the main re-establishment of the
curtain involves the ctenidial tips being ‘‘walked”’ (largely by
ciliary action) along the side and edge of the foot, and over
each other until an orderly row is again set up. With re-
establishment of the row, the ciliary junctions reconnect the
tip of one posterior leaflet either to one or to two anterior
leaflets on the gill behind it.

In healthy chitons, the way in which each ctenidial row
moves as a single dynamic curtain is impressive. It bulges
and flattens to accommodate changes in the hydraulics of the
pallial groove resulting from shifts in the inhalant (and less
frequently the exhalant) openings across the girdle as the
chiton crawls along. The early observation of Yonge (1939)
that inhalant openings can be formed by local lifting of the
girdle at almost any point along the anterior part of the chiton
is clearly confirmed. Yonge’s conjecture, that the capacity for
creating inhalant openings back along the sides of the body
is valuable when the anterior end is out of the water, can be
supported by the observation that, in Chaetop/leura at least,

RUSSELL-HUNTER: GILLS OF CHITONS 73

INA

Ve HW HiTT } )))

me y) 4h) )] WY

Wy! No

aaa Wy) | })

Fig. 3. Stereogram of part of a chiton ctenidium. Water is moved dorsally (and pedally) by bands of lateral cilia (1c). On the inhalant (INH)
side, the ctenidial leaflet edges bear front cilia (fc) and the ctenidial axis contains the efferent branchial vessel (ebv). On the exhalant (EXH)
side, the leaflet edges bear abfrontal cilia (ac) and the gill axis contains afferent branchial vessel (abv). The opposed free tips of the leaflets
bear specialized cilia forming the Velcro-like ciliary junctions (x), probably representing modified frontal cilia.

the continuity of the ctenidial curtain is maintained even with
air-bubbles in the anterior third of the groove on both inhalant
and exhalant sides of the gill row. Yonge (1939) also noted
that the exhalant opening across the girdle was less variable
in position, being always at the posterior end. At first sight
this seems true in living Chaetopleura, and the anus in the
midline is always swept by a strong exhalant current. However,
while the arching of the girdle to form an exhalant opening
always occurs close to the anus, its size (with water velocity
inversely related) and its direction (to left or to right of the
midline) do vary. As such changes occur, accommodation of
the ctenidial curtain to pressure shifts involves it becoming
less convex (more flattened towards the foot, decreasing the
exhalant cavity volume) or more convex (decreasing the in-
halant fraction of the pallial cavity). Changes resulting from
shifts in size or direction of the exhalant opening can be par-
ticularly obvious in a chiton crawling over a curved or irregular
surface. Once again, the simplest set-up used to view a chiton
through a flat glass surface can be deceptive.

Working with both living specimens and models of
mopaliid chitons, R. S. Cox and his colleagues have applied
water flow visualization techniques in flow tanks and have
noted muscular contractions of the pallial groove walls
(Douglas J. Eernisse, pers. comm.). They have had only
equivocal evidence of pallial shape producing augmentation
of flow (such as ramming or Bernoulli effects), but my obser-
vations suggest that the chiton’s ability to modify the exhalant

(downstream) pressure by changes in the effective diameter
of its exhalant girdle opening could have some significance
in shifting the fluid dynamics of the pallial system. Despite
this, basic water propulsion and consequent differential
pressures in the pallial compartments must all result from the
activity of the lateral cilia on the ctenidial leaflets. It is
noteworthy that, even in adult chitons, there are always some
bands of ciliated epithelia on the walls of the pallial groove
which beat in a posterior direction (particularly on the inside
of the girdle). Such ciliation is obvious in young (30-day)
postlarval chitons, where it exists before the first ctenidial buds
and creates analogous water currents (Russell Hunter and
Brown, 1965). However, in adults these cilia seem to propel
superficial strings of mucus rather than the ambient water.

Despite the adjustments of walls and openings, the
dynamic continuity of the ctenidial curtain is maintained as
the living chiton crawls along. The direction of the gill axes,
with their obvious efferent branchial vessels (Fig. 2), can be
seen to be altered but adjacent axes always stay more or less
parallel. Groups of six to eight (or occasionally more) gills
move together, with their gill-tips lagging behind the foot as
the chiton crawls forward, or making a fast recovery so that
the gill-tips are seen to be moving forward relative to the edge
of the foot. Similarly, groups of ctenidia acting together can
move their tips toward and away from the pedal edge. This
must involve neural coordination in, for example, the
simultaneous contraction of the afferent muscle strands in

74 AMER. MALAC. BULL. 6(1) 1988)

eight adjacent gills. Some part of the continuity of the cur-
tain could be passive after the ciliary junctions have been con-
nected, but there are obviously also active movements involv-
ing the coordination of several ctenidia or even most of the
ctenidial row. The ctenidial curtain sometimes shows a
metachronal wave of forward movement independent of the
foot, or a group of tips crawling together along the foot. Again
the loose or temporary attachment of ctenidial tips to the pedal
edge does not include any Velcro-like action, although
mucous-bound packages of cleansed material are often
passed to the foot. In addition, it was already noted that groups
of three or more gills could be simultaneously involved in the
faster cleansing reflex.

ADAPTIVE SIGNIFICANCE OF THE GILL CURTAIN

Perhaps the most important observation to be made
about the whole mantle groove system in chitons is that it is
dynamic. Unlike the pallial cavity of a bivalve or shelled
gastropod with its relatively static dimensions and shape, the
chiton pallial groove is a chamber bounded by pedal and gir-
dle walls whose shapes continually change with movements
of the chiton. The chamber wall provided by the habitat sur-
face (Fig. 1B) can also change markedly, since chitons can
and do crawl round corners and over edges. Thus the ctenidial
curtain has to conform (as a continuous, water-pumping,
porous partition) not only to the inhalant and exhalant imposed
pressure changes noted above but also to the shape changes
of the whole groove system. This is probably the reason why
chitons have evolved their pallial curtain by replication of a
series of gills rather than by the elongation of axes or of
filaments (leaflets) in one pair (or two pairs) of ctenidia. Leav-
ing aside consideration of the evolution of the higher
lamellibranch bivalves, the potential for hypertrophy of single
units of molluscan ctenidia is amply demonstrated in certain
gastropods. In Calyptraeid prosobranchs, a water-propulsive
ctenidial curtain is achieved by the elongation into filaments
of the leaflets of a single pectinibranch (one-sided) gill. It is
proposed that an adaptive functional explanation for the evolu-
tion of ctenidial replication in chitons is provided by the
dynamic nature of the mantle grooves in the group.

Admittedly, there are two obvious omissions in this
survey of the functioning of the ctenidial curtain in chitons.
First, there are almost no comparative data on gill function
in chitons with Lepidopleurid and other patterns of posterior
gills. Although many (perhaps most) species of chitons have
long gill rows essentially like those in Chaetopleura,
Lepidochitona and Tonicella, a variety of other conditions have
been described. Early workers, such as Pelseneer (1897),
developed a syntagma, or array of holobranch and mero-
branch forms, with metamacrobranchs and mesomacro-
branchs, and with or without adanales. A simpler, and pro-
bably more functionally significant, classification of certain
gill position characters has been utilized by D. J. Eernisse
(pers. comm.) in the course of revising the probable higher-
level phylogenetic relationships among chitons. Even the most
skeptical approach to the use of pallial cavity structures in
chiton classification has to separate the Lepidopleurids. Again
it would be helpful to know something of comparative gill func-

tion in these forms, as well as something of comparative
development (Minichev and Sirenko, 1984).

Recent studies on variation in larger population
samples of common European chitons (A. M. Jones, pers.
comm.) have emphasized the need for a population approach
to assessing taxonomic characters. Even in Chaetopleura,
usually described as holobranch, two distinct forms occur
within the populations studied at Woods Hole (Russell Hunter
and Brown, 1965) differing in the extent to which each pallial
groove is occupied by the ctenidial row. In one form the bases
of the gills extend forward for only about 75% of the pallial
groove, while in the other the bases extend anteriorly as far
as the head fold, and thus conform to the accepted species
diagnosis. It is possible that these could reflect phenotypic
growth responses to levels of microhabitat oxygenation, but
D. J. Eernisse (pers. comm.) has pointed out that, given the
lack of knowledge of these stocks, subsequent investigation
of other character states might well establish the two forms
as separate subspecies or even species. However, none of
the new observations presented in this paper would be in-
validated if it were subsequently proven that the studied
specimens of Chaetopleura apiculata from near Woods Hole
belonged in two distinct but congeneric species.

The second gap in this presentation on the function-
ing of the ctenidial curtain in chitons involves the lack of any
studies on the ultrastructure of the cilia concerned (particularly
those of the ciliary junctions). Any interested investigator with
access to SEM facilities, and appropriate techniques of nar-
cotization and fixation, could elucidate much of interest.

SUMMARY OF OBSERVATIONS

Even with these two major omissions, the observations
on gills in living chitons can be summarized as five topics.
First, the gills are not paired structures but can be added
asymmetrically during continued adult growth. Secondly, each
gill appears to be structurally and functionally homologous
with the aspidobranch ctenidium of archaeogastropods. Third-
ly, a neuromuscular cleansing reflex is common to the gills
of both chitons and archaeogastropods. Fourthly, each of the
two gill rows in chitons is organized as a coordinated ctenidial
curtain utilizing ciliary junctions. Fifthly, the adaptive
significance of ctenidial replication in chitons (rather than
hypertrophy of single units) could lie in the dynamic nature
of the pallial space.

DISCUSSION

Many aspects of the phylogeny of molluscs, and of
molluscan ancestry, remain controversial. The observations
presented here on the gills of living chitons have significance
only in relation to two of these aspects: first, the structural
and functional homologies of ctenidia and, secondly, the
possible metamerism of ancestral molluscs. They can con-
tribute little or nothing to other debates in molluscan
phylogeny, such as whether the primitive mantle-cavity was
a pallial groove surrounding the head-foot or a posterior cavity
with a complex of paired pallial structures, or if the primitive

RUSSELL-HUNTER: GILLS OF CHITONS 79

mantle was dome-shaped and secreted a one-piece shell.
Similarly, questions of the relationships between the three ma-
jor classes of ‘‘modern’”’ molluscs and the Aplacophora,
Monoplacophora and Polyplacophora are barely glossed by
this work. The two pertinent questions of ctenidial homologies
and of ancestral metamerism both merit further discussion,
but the former can be dealt with more simply and its near
enthymeme is set out first. Ancestral metamerism requires
both some conceptual history and more extensive and
multilateral exposition, and these will follow.

In evolutionary hypotheses, organ structures are con-
sidered homologous in two or more animal forms if they can
be claimed as being derived from a common precursor organ
structure in a common ancestral animal (Mayr, 1969, 1983;
Russell-Hunter, 1979). Such theoretical claims are normally
based on similarity of fundamental structural plan in the
organs concerned, on similar anatomical associations with
other organs, and on similarities in their embryonic develop-
ment. Since such claims are inferential, most modern evolu-
tionists would prefer them to be phrased in terms of maximum
likelihood. When, as in the case of the molluscan pallial cavity
and ctenidium, we have a whole concert of organs and func-
tions operating in an integrated fashion, there is likely to ex-
ist what can be termed functional homology (Russell-Hunter,
1968, 1979). It can be deduced that extensive patterns of func-
tional interdependence must be encoded by largish packets
of integrated genetic material commonly derived (since the
precursor animal must also have been an efficient machine
with similar functional interdependence). Cytogenetic levels
of linkage need not be postulated. On the other hand, attempts
at the enumeration of discrete unit characters for the
molluscan ctenidium and its associated pallial complex for
either cladistic (Hennig, 1950, 1966) or phenetic analysis
would be relatively uninformative from such an integrated
system (Mayr, 1974, 1983). The ctenidium, a gill with
characteristic patterns of ciliated epithelia and blood vessels,
is found as a homologous structure in Gastropoda, Bivalvia,
and Cephalopoda (Yonge, 1947).

In each mollusc with them, the ctenidia are part of an
integrated functional system: the heart and other blood
vessels, certain glands and sense-organs, the external open-
ings of genital and renal systems, and the posterior part of
the alimentary canal are all structurally and functionally
stereotyped in their relationships to the ctenidia. As pointed
out elsewhere (Russell-Hunter, 1968, 1979), it is highly signifi-
cant that, although probably at least 75,000 molluscan species
(out of about 110,000) have ctenidia, and although there are
many aquatic animals belonging to other phyla which seem-
ingly could make good use of a ctenidium, no nonmolluscan
animal has one. The above observations on gills in living
chitons can only confirm the conclusion reached by Yonge
(1939, 1947) that the gill rows represent multiplied ctenidia.
David R. Lindberg (pers. comm.) remains unconvinced of
homology between gills of chitons and those of gastropods,
largely on the basis of differences between the two classes
in the blood vessels draining the haemocoelic spaces of the
body and supplying the afferent branchial vessels of the gills.
However, it is not the preafferent circulation that links the

ctenidium with its associated pericardial and pallial structures
in a functionally homologous complex, but the postefferent
connections to the auricles, auriculoventricular openings, and
the rest that do so. Further, there is considerable variation
within gastropods in the arrangement of the preafferent
vessels. The attempt by Lemche (see especially Lemche,
1966) to suggest that bivalves and cephalopods have gills of
different origin from those of gastropods was based on a
misunderstanding of the relationships of their suspensory
ligaments in respect to the branchial vessels. It was associated
with his claims for homology between the gills of chitons and
those of Neopilina (Lemche, 1959a, 1966) and, in turn, be-
tween the gills of Neopilina and the limbs of arthropods like
trilobites (Lemche, 1959b, 1966). Each chiton gill is a true
ctenidium, structurally and functionally homologous with the
aspidobranch gills of Archaeogastropoda and the protobranch
gills of more primitive Bivalvia. Again, it has to be admitted
that this concluding hypothesis of homology for chiton gills
makes little contribution to the vexed questions of further
homology with the gills of Neopilina (Lemche, 1966) or with
the gills in certain Aplacophora (Scheltema, 1973, 1988).
The other phylogenetic controversy, that on metameric
segmentation in the ancestral mollusc, is less easy to set forth.
Before attempting to outline its history and arguments, some
statement of premises regarding both metameric segmenta-
tion and archetypes as models of ancestors may be ap-
propriate. The essence of metameric segmentation as found
in annelids and arthropods is the serial succession of
segments each containing unit-subdivisions of the several
organ systems (Hyman 1951; Russell-Hunter, 1968, 1979). The
sequence of morphogenesis of these segments is antero-
posterior from a penultimate budding zone, so that the
segments just behind the head are older, and the more
posterior ones (just in front of the budding zone) are younger.
The differentiation of additional segments in this mode of
morphogenesis is such that each new segment contains (at
least initially or potentially) a full set of all organ systems.
Archetypes are not ancestors. For any stock of animals,
the characteristics of the actual ancestral forms will never be
known with certainty. Archetypes are logical constructs, tem-
porary models set up from reductionist explanations of
available data, to be tested by the collection of further data.
The testing can invalidate, but can never authenticate (despite
the current belief of certain systematists that their cladistic
hypotheses can be confirmed by separately computed
phenetic analyses). When considering such models, in view
of what Mayr (1983) terms ‘‘cohesion of the genotype’’, it
seems particularly important to consider possible functional
homologies as well as the more usual morphological ones
(Russell-Hunter, 1979). Significant functional unity is apparent
within each phylum of more complex animals (including
molluscs, arthropods, echinoderms, and chordates). There
are obvious pragmatic values in setting up ancestral models.
There are peculiar dangers in evolutionary discussions after
setting up an archetype, and these seem to result from
assembling together in the unfortunate hypothetical animal
a group of incompatible structures, all thought to be
‘primitive’ or ‘‘plesiomorphic’’ within the stock. As noted

76 AMER. MALAC. BULL. 6(1) (1988)

elsewhere (Russell-Hunter, 1968, 1979), many of these
dangers can be avoided if, when a hypothetical ancestral type
is constructed, an attempt is made to create a working arche-
type — one in which the concert of organs and functions could
operate as a whole, in an integrated functional plan, as in all
living organisms. In discussing similar matters in the adap-
tive morphology of vertebrates, Bock (1965) (see also Bock
and von Wahlert, 1965) has clearly stated the need for
analyses of function in the whole animal. The working arche-
type (Russell-Hunter, 1968, 1979) can be set up from a de-
duced concert of structures and functions together forming
an integrated functional plan, and can then provide a better
basis for phylogenetic speculation and both predictive and
retrodictive testing.

Molluscan archetypes with short segmented bodies
had been proposed by Pelseneer (1899, 1906) and Naef (1926),
largely on the basis of studies on the genital and excretory
systems of chitons and cephalopods. However, the extensive
and convincing work of the molluscan functional morpho-
logists such as Yonge, Graham and Fretter on ciliary
mechanisms, ctenidial blood vessels, and renopericardial and
genital ducts (particularly in more primitive gastropods) set
up a very different model for the stem-mollusc. As set out in
fecund summary by Yonge (1947), although primitively
bilaterally symmetrical, this archetype was totally unseg-
mented and possessed a posterior mantle-cavity enclosing
a pallial complex of paired structures which included two
ctenidia. This model convincingly survived retrodictive testing
against the fossil record, as clearly set out by Knight (1952)
who was able to fit appropriate pallial circulation and muscle
attachments into the lower palaeozoic monoplacophoran
genera, Scenella and Pilina, regarded then as untorted
“‘pregastropods.”’ Pragmatically, it is important to note that
versions of Yonge’s model are still employed in the 1980’s by
systematists (Salvini-Plawen, 1980; Seed, 1983) and
pedagogues (Russell-Hunter, 1979, 1982) both as gastropod
archetype and as bivalve archetype and, as regards the paired
pallial structures and homologous ctenidia of these two stocks,
have survived much testing.

Discussion of possible metamerism in ancestral
molluscs was reopened by the discovery of a living monopla-
cophoran, Neopilina, by its preliminary description (Lemche,
1957) and by the extensive description of its morphology
(Lemche and Wingstrand, 1959) that followed. It was hypothe-
sized that the mollusc ancestor must have shown relatively
complete metamerism, that this is present to a somewhat
reduced extent in Neopilina, that this is still further reduced
in chitons, and that this metamerism degenerates so com-
pletely as to be undetectable in gastropods and bivalves
(Lemche and Wingstrand, 1959). Subsequently Lemche (1966)
reversed part of this hypothesis and claimed that the arthro-
pods originated directly from a molluscan ancestor. For a few
years, many strange phylogenies were based on Neopilina
as a ‘‘missing link’’ rather than as an interesting survivor of
a less successful molluscan stock. In this respect, the claims
of homology among the gills of chitons, the gills of Neopilina,
and the arthropod limbs of trilobites (Lemche 1959b, 1966)
begin to approach the idealist metabiological comparisons

of William Patten. As noted elsewhere (Russell-Hunter, 1985),
Patten’s use, early in this century of detailed comparative
anatomy to postulate an origin of vertebrates in arachnids (or
merostomatids like Limulus), represents a comparatively late
derivative of the Naturphilosophen of Johann Wolfgang von
Goethe (1749-1832), and is perhaps closest in concept to the
publications of Lorenz Oken in the first half of the nineteenth
century. Even without idealist morphology, in the work of
Lemche and Wingstrand (1959) on Neopilina, and in the
beautiful reconstructions subsequently presented by
Wingstrand (1985), it is explicit that the multiplied organs of
chitons (shells-valves, muscles, gills and nerves) reflect
metameric segmentation. Indeed, after detailed comparisons
of Neopilina, Vema and chitons, Wingstrand (1985) concludes
that a homologous 8-metamerism is present in the Poly-
placophora. Such a chiton archetype with true metamerism
can be tested appropriately with the data on actual replicated
structures in chitons including the numbers and symmetry
of gills (Russell Hunter and Brown, 1965), and the function-
ing of the gill series (this paper). Even when the other
multiplied structures are considered, there is little of the serial
succession of segments, each with unit subdivisions of organ
systems, in any living chiton, and there is no evidence of serial
organogenesis. The mantle rudiment of a settled postlarval
chiton secretes six plates. After an interval a larger anterior
plate is added then, still later, a small posterior plate. There
is never a budding zone as in the annelid-arthropod mode
of development. Segmentation in heart structures is even less
valid. Chitons all have an elongate ventricle in the midline
which receives blood from two symmetrical elongate auricles.
Most chitons have two pairs of auriculoventricular openings,
several genera have one pair, and chiton species are known
with three pairs and with four pairs. Both Neopilina and
Nautilus have four auricles and therefore also have two pairs
of auriculoventricular openings. Individual ctenidia in chitons
cannot be related to any other replicated organs, such as shell-
valves, nephridial lobes, lateropedal nerve connections or
heart structures, and thus cannot be allocated to specific
metameric segments. Other features of chiton gills and their
functioning complete this negation of the metameric archetype
for chitons. The gills are not paired but are added asym-
metrically during continued adult growth. As individual gills,
they seem to be structurally and functionally homologous with
those of primitive bivalves. This replication in chitons results
in gill rows, which show coordinated function as pallial cur-
tains and cannot reflect simplification of more extensive
metamerism. Ctenidial replication in chitons can be claimed
to result adaptively from the dynamic nature of the pallial
grooves in the chiton body form.

Similar arguments can be used to criticize the concept
of annelid-arthropod metamerism applied to the described
structures of Neopilina and Vema. This statement should not
be taken as critical of the majority of the interesting
homologies elucidated by Wingstrand (1985), in particular his
meticulously exhibited parallels between chitons and the two
monoplacophoran genera not only in pedal retractor muscles
but also in the muscles of the buccal mass and radula.
However, living monoplacophorans have five (or six) pairs of

RUSSELL-HUNTER: GILLS OF CHITONS 77

gills, eight pairs of pedal retractor muscles, two pairs of
auricles, six (or seven) pairs of nephridiopores, two (or three)
pairs of gonads, and a single shell (Wingstrand, 1985). This
assemblage is unlikely to have arisen by segmental morpho-
genesis.

In his claims for molluscan metamerism, Wingstrand
(1985) appears to rely on the concept of a monophyletic Pro-
tostomia or Spiralia, linked by common features of early
cleavage, gut development and larval type. It may be best to
quote his own words (Wingstrand, 1985: 89): ‘“‘The
metamerism of molluscs is in itself hardly unexpected, for
many features support their incorporation within the Spiralia,
a group in which different kinds of metameric repetition are
common.” Unfortunately, the concept of a group of phyla form-
ing the Spiralia is itself suspect. The five diagnostic features
used to discriminate the group from the Deuterostomia are
neither so universal nor so consistent as to justify a clear
dichotomy (Russell-Hunter, 1979). Larval homologies have
been in doubt since Garstang (1922, 1929) seriously chal-
lenged recapitulation as an important factor in the evolution
of larval stages. Cleavage is a dynamic process in time and
spiral cleavage is not absolutely correlated with mosaic
development. As Costello (1948, 1955) pointed out, there are
three main categories of cleavage (radial, bilateral and spiral),
and three basic types of spiral cleavage (by quartets, by duets
and by monets), but all are modified into bilateral cleavage
later in development. He emphasized that the occurrence of
spiral cleavage has no obvious significance in the interrela-
tionships of animal phyla (Costello and Henley, 1976).

There is another kind of developmental evidence link-
ing molluscs and flatworms and making molluscan
metamerism less likely. Recent work on actuarial bioener-
getics has emphasized the capacity for degrowth in some
shelled molluscs (Russell-Hunter et a/., 1983, 1984; Russell-
Hunter, 1985), and compared it in flatworms. Along with other
features of indeterminate growth, many gastropods and
bivalves show a capacity to degrow (as individuals to reduce
the mass of their structural proteins under certain circum-
stances), no close-coupling of growth with sexual maturation,
and a lack of endogenous senescence (Russell-Hunter and
Eversole, 1976; Russell-Hunter and Buckley, 1983; Russell-
Hunter, 1985). It has been hypothesized (Russell-Hunter, 1985)
that this capacity in flatworms and molluscs could involve con-
trols of genetic expression that cannot coexist with those in-
volved in a metameric pattern of morphogenesis. Some
molecular biologists studying ageing indicate accumulated
errors in the synthesis of macromolecules as important
(Kirkwood, 1977; Kirkwood and Holliday, 1979), and they cor-
relate the absence of endogenous senescence in certain
organisms with indeterminate growth patterns. The neuro-
hormonal and hormonal controls for metameric development
may mandate selective gene-expression in some irrevocable
fashion that is incompatible with cellular dedifferentiation-
rejuvenation, and with the capacity of degrowth exhibited by
molluscs and flatworms. This hypothesis of incongruent con-
trols of morphogenesis in molluscs and in metamerically
segmented animals cannot yet be tested experimentally. That
it can be proposed illustrates the weight of circumstantial

evidence that metameric organogenesis of the sort which pro-
duces serial sets of structures in the phyla Annelida and
Arthropoda never occurs in the Mollusca.

As already admitted, conditions in the stem-mollusc
remain controversial. The general conclusion from the pre-
sent work that chitons do not show true metameric segmen-
tation seems established at a high level of likelinood. Extend-
ing the logic, evidences against metamerism of the annelid-
arthropod pattern in all primitive molluscs are strong, and a
consensus with the views of Wingstrand and of Salvini-Plawen
could be achieved if their protoannelid ancestor for the
molluscan stock were totally without metameric segmenta-
tion, indeed if it were an unsegmented flatworm turned
coelomate. All model ancestors are highly speculative.

At the end of the earlier paper on chiton gills (Russell
Hunter and Brown, 1965), an archetype mollusc with a four-
fold basic organization (that is, with four ctenidia, four auricles,
four renal organs, etc.) was proposed. This derived from a
footnote query by C. F. A. Pantin in Yonge (1947), and reflected
the heart morphology of Neopilina, Nautilus and chitons.
Somewhat surprisingly, this model is mentioned favorably not
only by Minichev and Sirenko (1984) but also in passing by
Wingstrand (1985). From such a four-fold organization, two
sorts of subsequent morphogenesis could occur. Both a line
of organisms with one gill on either side, and a line with many,
could thus evolve from an archetype with two pairs of gills.
In this hypothesis, the former stock (that is, those with one
pair of ctenidia, one pair of auricles, one pair of renal organs,
and so on) could still be regarded as archetypic for the two
major groups of living molluscs: the gastropods and the
bivalves. But, as reiterated pedantically here and elsewhere,
archetypes are not ancestors.

ACKNOWLEDGMENTS

Along with my sincere thanks to Robert C. Bullock for organiz-
ing the polyplacophoran symposium at Key West, must also go thanks
to William G. Lyons for initiating it, and to my fellow participants for
profitable discussions. My wife, Myra Russell-Hunter, once again aided
in the production of this paper and, at intervals over 26 years, helped
me collect chitons. | must also thank John J. Valois, head of the marine
resources department of the Marine Biological Laboratory, Woods
Hole, who added to earlier help by providing at short notice working
space for two days (and a few chitons) in the Spring of 1987. Some
matters considered here reflect long dormant ideas seeded in discus-
sions with two colleagues 24 and 27 years ago. The work is a belated
continuation of chiton gill-counting and phyletic talk around 1963 with
Stephen C. Brown, now of the State of University of New York at
Albany. It also benefits from debates with my dialectical colleague
in the University of Glasgow, Roy A. Crowson, an early and commit-
ted cladist. Of course, the paper only adds a few bricks to the foun-
dations laid down by the late C. M. Yonge, who was my undergraduate
research advisor more than forty years ago. | am grateful to all three.

| must also thank Barbara J. Carns for help in the preparation
of the manuscript. Partial support for earlier work came from research
grant DEB-78-10190 from the National Science Foundation and for
the final months from a Senate Research Award from Syracuse
University.

78 AMER. MALAC. BULL. 6(1) (1988)

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Date of manuscript acceptance: 24 November 1987

A REVIEW OF CARIBBEAN ACANTHOCHITONIDAE (MOLLUSCA:
POLYPLACOPHORA) WITH DESCRIPTIONS OF SIX NEW SPECIES
OF ACANTHOCHITONA GRAY, 1821

WILLIAM G. LYONS
FLORIDA DEPARTMENT OF NATURAL RESOURCES
ST. PETERSBURG, FLORIDA 33701, U. S. A.

ABSTRACT

Nine previously described species of Acanthochitonidae are recognized in the region between
Bermuda and the Caribbean coast of South America: Acanthochitona andersoni Watters, 1981; A.
astrigera (Reeve, 1847); A. balesae Abbott, 1954 (+A. elongata and A. interfissa, both Kaas, 1972); A.
bonairensis Kaas, 1972; A. hemphilli (Pilsbry, 1893); A. pygmaea (Pilsbry, 1893); A. rhodea (Pilsbry, 1893);
Choneplax lata (Guilding, 1829); Cryptoconchus floridanus (Dall, 1889). Four new species (Acanthochitona
lineata, A. roseojugum, A. worsfoldi, and A. zebra) are described from Florida, the Bahama Islands
and the northern Caribbean; Acanthochitona venezuelana sp. nov. is described from Margarita Id.,
Venezuela; Acanthochitona ferreirai sp. nov. is described from Pacific coasts of Panama and Costa Rica.
No subsequently collected specimens were seen of Acanthochitona spiculosa (Reeve, 1847), original-
ly described from the West Indies; A. spiculosa is considered a species inquirenda.

Until recently, seven species of Acanthochitonidae
generally were recognized in the Caribbean region (Bermuda,
Florida, and the Bahama Islands to the north coast of South
America): Acanthochitona spiculosa (Reeve, 1847) [+A.
astriger (Reeve, 1847)]; A. hemphilli (Pilsbry, 1893); A.
pygmaea (Pilsbry, 1893); A. rhodea (Pilsbry, 1893); A. balesae
‘Pilsbry’ Abbott, 1954; Choneplax lata (Guilding, 1829); Cryp-
toconchus floridanus (Dall, 1889). In 1972, P. Kaas: published
a monograph on the Polyplacophora of the Caribbean region.
In his treatment of Cryptoplacidae (=Acanthochitonidae), Kaas
(1972) proposed A. elongata to replace A. balesae ‘Pilsbry’,
recognized as valid the other six species, and described two
new species, A. bonairensis and A. interfissa, increasing to
nine the number recognized from the region. Kaas was fol-
lowed by G. T. Watters’ (1981) review of New World Acantho-
chitona, in which he declared A. astriger to be separate from
A. spiculosa, assigned A. pygmaea to the synonymy of A.
spiculosa, assigned A. rhodea to the synonymy of A. hem-
philli, resurrected A. balesae Abbott, with synonyms A.
elongata and A. interfissa, declared A. bonairensis to be a
synonym of the European A. communis (Risso, 1826), and
described a new species, A. andersoni. As a result, the
number of recognized Caribbean species of Acanthochiton-
idae was reduced to eight.

In a report on the Polyplacophora of Barbados pub-
lished four years later, A. J. Ferreira (1985) proposed addi-

tional changes in the classification of Caribbean Acanthochi-
tonidae. Ferreira recognized Acanthochitona astrigera, A.
spiculosa, A. bonairensis, and Cryptoconchus floridanus,
reversed Watters’ action by assigning A. hemphilli to the
synonymy of A. rhodea, and declared A. andersoni, A. balesae,
and A. interfissa to be juveniles, and thus synonyms, of
Choneplax lata. Six recognized species of Caribbean Acan-
thochitonidae remained.

In this report, | present new conclusions based upon
examination of type specimens of Acanthochitona andersoni,
A. astrigera, A. bonairensis, A. hemphilli, A. interfissa, A.
pygmaea, A. rhodea, and A. spiculosa. | have relied exten-
sively on specimens in the collection of the Florida Depart-
ment of Natural Resources (FDNR) and in the research col-
lection of Dr. R. C. Bullock, University of Rhode Island. | also
re-examined many museum specimens utilized previously by
Kaas (1972), Watters (1981), and Ferreira (1985). After Dr.
Ferreira’s death in 1986, his collection was transferred to the
California Academy of Sciences. Unfortunately, only the dry
collection could be inspected during 1987.

Nine previously named species and five new species
of Acanthochitonidae that occur in the Caribbean region are
described and illustrated. Relationships between Caribbean
species and their eastern Pacific cognate species are dis-
cussed, and one eastern Pacific cognate species is described
as new.

American Malacological Bulletin, Vol. 6(1) (1988):79-114

79

80 AMER. MALAC. BULL. 6(1) (1988)

METHODS

Complete species treatments should provide full
descriptions and illustrations of valves, spicules and radulae.
Examinations of spicules and radulae of taxa treated here are
still in progress and so cannot be presented. Instead, this
report presents conclusions derived principally from
characters of the valves, with less emphasis on girdle spicules
and no information on radulae. Descriptions, illustrations, and
differential diagnostic comments are provided for all of the
species. Characters described include general dimensions,
color, shape of the jugum, tegmentum, sutural laminae, and
insertion plates, tegmental pustule morphology, and counts
and measurements of girdle spicules. Species are illustrated
with SEM photographs of valves and tegmental pustules as
well as with photographs of intact specimens.

Tegmental pustules are illustrated in near-perpendicu-
lar aspect from anterolateral portions of left or right sides of
intermediate valves, depending upon specimen condition. As
shown in illustrations of entire valves, pustules usually are
arranged in rows parallel to the jugum, but individual pustules
are aligned anterolaterally, so pustular apices point postero-
laterally toward the jugum.

Longitudinal lines or incisions on the jugum are men-
tioned often in descriptions of Acanthochitona. In fact, lines
are visible within the jugum of nearly all species examined,
but lines at the surface are uncommon. Careful examination
in most instances reveals that such lines are internal and do
not interrupt the jugal surface. Whether the surface is smooth
or incised can be ascertained by using scanning electron
microscopy or, with light microscopy, by using high magnifica-
tion with light directed obliquely at a low angle across the short
axis of the jugum.

Measurements of small intact specimens, individual
valves, and girdle spicules were made using a Zeiss IV-B
dissecting microscope with ocular micrometer. Dimensions
of tegmental pustules were measured from scanning electron
micrographs of known magnification. Large intact specimens
were measured with vernier calipers. Most specimens were
flat when preserved, so measurements are accurate to
0.1 mm. Lengths of slightly curled specimens were determined
by making several incremental linear measurements along
the longitudinal curve; those lengths are accurate to about
0.5 mm. Extremely curled specimens were not measured.
Data presented for individual species lots include number of
specimens, size range (total length), location, depth, date of
collection, and museum catalogue number.

Specimens were examined from or deposited in the
following institutional collections: Academy of Natural
Sciences of Philadelphia, Pennsylvania (ANSP); British
Museum (Natural History), London [BM(NH)]; California
Academy of Sciences, San Francisco (CAS); Delaware
Museum of Natural History, Wilmington (DMNH); Florida
Department of Natural Resources, Bureau of Marine
Research, St. Petersburg (FSBC 1); Indian River Coastal Zone
Museum, Harbor Branch Oceanographic Institution, Ft.
Pierce, Florida (IRCZM); Rijksmuseum van Natuurlijke
Historie, Leiden (RMNH); Tulane University Department of

Geology, New Orleans (TUDG); and the National Museum of
Natural History, Smithsonian Institution, Washington, D. C.
(USNM).

SYSTEMATIC ACCOUNTS

Family Acanthochitonidae Pilsbry, 1893
Genus Acanthochitona Gray, 1821

Acanthochitona hemphilli (Pilsbry, 1893)
Figs. 1-9

Acanthochites (Notoplax) hemphilli Pilsbry, 1893: 34, 35, pl. 13,
figs. 65-67.

Acanthochitona hemphilli, Kaas, 1972: 38-41, figs. 58-64, pl. 2,
figs. 1, 2 (pars), Watters, 1981: 173 (pars).

Acanthochitona rhodea, Ferreira, 1985: 207, 208 (pars) [non
A. rhodea (Pilsbry, 1893)].

TYPE MATERIAL: LECTOTYPE: = 24 mm, partially disarticulated;
Key West; ANSP 35803 (herein designated).

OTHER MATERIAL EXAMINED: FLORIDA: 3 spec., 33.4-
34.5 mm, Long Key Reef, Dry Tortugas, intertidal, 11-12 May 1979,
FSBC | 32042. —1 spec., 13.0 mm, patch reef near Long Key Reef,
1.5-2.5 m, 11 May 1979, FSBC | 32428. —2 spec., 33.5, 38.4 mm, Bird
Key Reef, Dry Tortugas, 0.5-1.0 m, 4 Oct 1979, FSBC | 32044. —8 spec.,
23.8-44.2 mm, Garden Key, Dry Tortugas, 1-2 m, 13 May 1979, FSBC
| 32043. —6 spec., 15.2-44.1 mm, Garden Key, 0-2 m, 5 Oct 1979,
FSBC | 32045. —1 spec., 29.3 mm, Sand Key off Key West, 0.5-2.0
m, 3 Aug 1980, FSBC | 32046. —2 spec., 30.0-36.2 mm, Western
Sambo Reef off Key West, 4.2-7.3 m, 12-21 Mar 1973, FSBC | 9397.
—1 spec., 50.9 mm, off Pompano Beach, southeast Florida, 18.3 m,
1981, FSBC | 32429. BAHAMAS: 27 spec., 10.0-51.3 mm, Bahama
Beach Canal, West End, Grand Bahama, intertidal, 29 Aug 1984,
FSBC | 32049. —5 valves, Gold Rock, Grand Bahama, bottom
sediments, 24.4 m, May-July 1981, FSBC | 32519. —14 spec., 8.0-
37.5 mm, McLeanstown, east end Grand Bahama, 1-2 m, 24 May 1981,
FSBC | 32047. —19 spec., 4.5-47.0 mm, McLeanstown, 1 m, 27 Aug
1984, FSBC | 32048. —8 valves, Grand Bahama, bottom sediments,
May 1981, R. Quigley collection. —11 spec., 30.5-47.6 mm, Harbour
Id., Eleuthera, 0-3 m, 24 Aug 1978, FSBC | 32041. —3 spec., 23.8-
41.9 mm, Fernandez Bay, Cat Island, 3 m, 10-16 July 1976, FSBC |
15804. —1 spec., curled, Georgetown, Great Exuma, 0-1 m, 21 June
1974, FSBC | 32518. TURKS AND CAICOS ISLANDS: 8 spec., 11.1-
51.3 mm, Providenciales, 0-2 m, 22 Sept 1986, FSBC | 32430.
PUERTO RICO: 2 spec., 38.8, 41.6 mm, Cayo Enrique, La Parguera,
0-1 m, 19 Aug 1985, FSBC | 32050. —8 spec., 15.2-32.1 mm,
Magueyes ld., La Parguera, 20 Apr 1966, Bullock collection. JAMAICA:
1 spec., 30.8 mm, 3 km west of Runaway Bay, 0-1.5 m, 3 Nov 1983,
Bullock collection. CAYMAN ISLANDS: 1 spec., 35.8 mm, Grand
Cayman, 1965, FSBC | 5549. BELIZE: 2 spec., curled, Carrie Bow
Cay, 8 m, 21-24 Oct 1973, FSBC | 10765. —8 spec., curled, Carrie
Bow Cay, 23 Mar 1981, IRCZM 61:050. HONDURAS: 2 spec., 22.6,
34.5 mm, Anthonys Key, Roatan, 4-10 July 1971, Bullock collection.
—13 spec., all curled, Oak Ridge, Roatan, intertidal, Mar 1987, FSBC
| 32431. —5 spec., 24.0-32.1 mm, Roatan, 1981, FSBC | 32520.

TYPE LOCALITY: Key West, Florida (original designation).

DISTRIBUTION: South Florida and Grand Bahama Island to
Puerto Rico, Jamaica, and Honduras; intertidal to 18 m.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 81

Figs. 1-6. Acanthochitona hemphilli (Pilsbry, 1893). Fig. 1. Whole specimen, 42.6 mm; Harbour Id., Eleuthera, Bahamas; FSBC | 32041.
Fig. 2. Valve i ex 15.2 mm specimen; Dry Tortugas, Florida; FSBC | 32045. Fig. 3. Valve iv, same specimen. Fig. 4. Valve viii, same specimen.
Fig. 5. Tegmental pustules, valve v, 16.7 mm specimen; same lot (field width = 330 um). Fig. 6. Spicules of dorsal girdle mat, specimen from

McLeanstown, Grand Bahama; FSBC | 32047 (field width = 500 um).

DESCRIPTION: Largest specimen 51.3 mm long, 28.0 mm
wide including girdle; valves occupying about 30% of total
specimen width (Fig. 1). Exposed parts of valves dark red with
white maculations, small relative to total specimen size; unex-
posed valve parts greenish white. Girdle broad, fleshy, ap-
pearing smooth, dark brown, with few brown or reddish brown
spicules in dorsal tufts; color of girdle and tufts sometimes
faded in preserved material.

Valve i semilunate (Fig. 2), wider than long, beaked,
sinuous posteriorly, with anterior insertion plate bearing 5 slits;
tegmentum occupying about 65% total valve length. Valves
ii-vii strongly beaked (Fig. 3); tegmentum spade-shaped, about
as long as wide, strongly constricted anteriorly, with broadly
sinuous anterolateral margins; sutural laminae broad, expand-
ed anteriorly, with subparallel lateral margins rendering overall
valve shape nearly quadrate except for broad, shallow anterior
sinus; single small narrow slits near midpoints of margins.
Valve viii subtriangular (Fig. 4), rounded posteriorly, with
elevated mucro posterior of center; tegmentum drop-shaped,
longer than wide, constricted anteriorly; sutural laminae large,
flared anterolaterally, with straight to sinuous anterior margins,
separated by wide V-shaped sinus; 2 slits in posterior inser-

tion plate small, narrow, V-shaped.

Jugum smooth, narrow, with parallel sides well-
separated from lateral tegmental surface. Tegmentum of all
valves covered with small (35-50 »m), round to ovate, cupped
pustules with edges incised at apex to render overall ap-
pearance reniform (kidney-shaped) (Fig. 5), with single cen-
tral macresthete, 2-3 micresthetes.

Girdle upper surface appearing smooth, actually
covered with dense mat of very small (50 »m) slender, sharp,
brown spicules (Fig. 6); 18 anterior and sutural dorsal tufts
with about 50 thick, reddish-brown to white, flat-sided spicules,
longest 1.5-2.0 mm; slender, needle-like, sharp spicules in-
terspersed among larger spicules of tufts; margin with dense
fringe of straight, slender, brown and white spicules 1.0-1.5
mm long; underside covered with small (50 »m) slender, sharp,
white spicules directed toward periphery.

DISCUSSION: Acanthochites hemphilli Pilsbry, 1893, was de-
scribed from a specimen collected at Key West, Florida, and
the name generally has been applied to the large
(>50 mm) fleshy species that occurs in south Florida, the
Bahama Islands, and the northern Caribbean Sea. A. rhodeus

82 AMER. MALAC. BULL. 6(1) (1988)

Pilsbry, 1893, was described from a specimen bearing only
the data ‘‘Panama (McNeill Expedition)’, prompting subse-
quent question as to whether A. rhodea properly belonged
to the Caribbean fauna, the eastern Pacific fauna, or perhaps
to both. Leloup (1941) reported specimens of Acanthochiton
rhodeus from off Cabo la Vela, Caribbean Colombia, and pro-
vided additional descriptive notes for the species. Keen (1958)
listed Acanthochitona rhodea in her compendium of mollusks
from the eastern Pacific but noted that the species might
belong to the Caribbean rather than the Panamic fauna.
A. G. Smith (1961) seemed to confirm the presence of A.
rhodea in the eastern Pacific when he contrasted its characters
with those of A. tabogensis Smith, 1961, and A. hirundiniformis
(=hirudiniformis) (Sowerby, 1832), two other species from that
region. The name again appeared on a list of Caribbean fauna
in Houbrick’s (1968) account of species from Costa Rica.
Thorpe (/n Keen, 1971) illustrated a specimen identified as A.
rhodea and listed its range as Mexico (Pacific Ocean) to Peru.
Kaas (1972) summarized descriptions by Pilsbry and by Leloup
and treated the species as a member of the Caribbean fauna.
The species again was illustrated and reported from Carib-
bean Colombia by Gotting (1973). Watters (1981) relegated A.
rhodea to the synonymy of A. hemphilli without discussion of
morphological characters or geographic range, only to be
followed soon thereafter by Ferreira (1985) who declared A.
hemphilli to be a synonym of A. rhodea, citing page priority
of the original descriptions. Ferreira concluded that the com-
plex constituted a single species ranging from Florida and the
Bahamas to Brazil in the western Atlantic Ocean and from Mex-
ico to Peru in the eastern Pacific Ocean. Examination of type-
specimens of both species, as well as additional materials
from Florida, the Bahama Islands, several localities in the
northern Caribbean, the Caribbean coast of Central America,
and the Pacific coasts of Costa Rica and Panama, indicates
that the complex actually consists of three species, including
one previously undescribed.

One of Pilsbry’s specimens (ANSP 35803) is herein
designated as lectotype of Acanthochites hemphilli Pilsbry,
1893. Pilsbry described a dried specimen 24 mm in length.
The lectotype measures about 24 mm overall and is partially
disarticulated (Fig. 7); valves i-vi remain attached to the gir-
dle, but valves vii and viii are free. Pilsbry described the
posterior valve viii as ‘...not bilobed behind, having the usual
two slits, and between them a number (6-8) of smaller, ir-
regular and unequal slits or nicks’’. That this is the specimen
described by Pilsbry is confirmed by the condition of valve
vill, which is aberrant. The valve has two slits in the usual
positions (Fig. 8), but one slit is unusually large and wide,
whereas the other is unusually small and narrow; the reported
irregularities are also present.

Asymmetrical tail valves are not unusual in Acantho-
chitona; several valves viii of A. astrigera which | examined
were misshapen, some completely lacking one of the posterior
slits. All other characters of the lectotype, including the
reniform tegmental pustules (Fig. 9), indicate the specimen
to be conspecific with material reported here as A. hemphilli.
The reniform pustules, “‘smooth’’ girdle, greenish white sutural
laminae, and subquadrate, parallel-sided intermediate valves

Figs. 7-9. Acanthochitona hemphilli (Pilsbry, 1893), lectotype; Key
West, Florida; ANSP 35083. Fig. 7. Whole specimen. Fig. 8. Valve
viii, ventral. Fig. 9. Tegmental pustules, valve vii (field width =
345 um).

distinguish A. hemphilli from A. rhodea and from the new
species.

Acanthochitona hemphilli now is demonstrated to oc-
cur from southeast Florida and the northern Bahama Islands
southward to Puerto Rico, Jamaica, Belize, and Honduras.
Specimens reported from Cuba (Jaume and Sarasua, 1943)
and Caribbean Mexico (Vokes and Vokes, 1983) are probably
referable to this species, whereas those reported from Carib-
bean Panama (Olsson and McGinty, 1958) almost certainly
are A. rhodea (see that species account). Specimens il-
lustrated by Kaas (1972: pl. 2, figs. 1, 2) from Curagao as A.
hemphilli are A. rhodea. Other reports of A. hemphilli from Bar-
bados, Bonaire, and Venezuela (Ferreira, 1985) and Aruba
(Kaas, 1972) could also represent A. rhodea. Records by Righi
(1971) of A. hemphilli in Brazil seem especially unlikely

LYONS: CARIBBEAN ACANTHOCHITONIDAE 83

because the specimens were collected in 47-115 m depths,
far deeper than the 18 m maximum depth otherwise known
for the species.

Acanthochitona rhodea (Pilsbry, 1893)
Figs. 10-18

Acanthochites rhodeus Pilsbry, 1893: 26, 27, pl. 12, figs. 48-51.

Acanthochiton rhodeus, Leloup, 1941: 39-42, figs. 5-7.

Acanthochitona rhodea, Keen, 1958: 519 (pars). Kaas, 1972:
42, 43, figs. 65-71. Gotting, 1973: 251-253, pl. 11, figs.
15, 16. Bullock, 1974: 164 (pars). Ferreira, 1985: 207,
208 (pars).

Acanthochitona rhodeus, Houbrick, 1968: 10, 20.

Acanthochitona hemphilli, Kaas, 1972: pl. 2, figs. 1, 2 (pars).
Watters, 1981: 173 (pars) [non A. hemphilli (Pilsbry)].

TYPE MATERIAL: HOLOTYPE: 3 disarticulated valves, ‘Panama;
McNeill Exped.’, ANSP 63429.

OTHER MATERIAL EXAMINED: COSTA RICA: 2 spec., both
curled, Portete, Limon Prov., 12 June 1966, USNM 702874. PANAMA:
1 spec., curled, Toro Point, Ft. Sherman, Canal Zone, Sept 1969,
Bullock collection. —4 spec., 6.5-27.0 mm, Ft. Randolph, Canal Zone,
1m, Nov 1980, FSBC | 32530. —10 spec., 1.8-30.0 mm, Galeta Id.,
Canal Zone, Bullock collection. —20 spec., 19.7-32.5 mm, Galeta Id.,
Sept 1973, FSBC | 32562 (1), Bullock collection (19). —2 spec., 10.3,
25.1 mm, near Portobelo, 0-1 m, Nov 1980, FSBC | 32529. —2 spec.
12.0, 12.4 mm, Cocal Point, Portobelo, 13 Sept 1973, Bullock collec-
tion. —12 spec., 24.5-39.5 mm, Ironcastle Point, Portobelo, 13 Sept
1973, Bullock collection.

TYPE LOCALITY: Portobelo, Caribbean coast of Panama (by
subsequent designation, Ferreira, 1985).

DISTRIBUTION: Caribbean coasts of Costa Rica, Panama,
and Colombia; intertidal to 53 m.

DESCRIPTION: Largest specimen slightly curled, 39.5 mm
long, 24.0 mm wide including girdle; valves occupying approx-
imately 30% of total specimen width (Fig. 10). Exposed parts

Figs. 10-15. Acanthochitona rhodea (Pilsbry, 1893). Fig. 10. Whole specimen, 25.1 mm; Portobelo, Panama; FSBC | 32529. Fig. 11. Valve i
ex 25.0 mm specimen; Galeta Id., Panama; FSBC | 32562. Fig. 12. Valve iv, same specimen. Fig. 13. Valve viii, same specimen. Fig. 14.
Tegmental pustules, valve iv, same specimen (field width = 340 ym). Fig. 15. Spicule clusters, dorsal girdle mat, same specimen (field width

= 550 pm).

84 AMER. MALAC. BULL. 6(1) (1988)

of valves dark red with white maculations; unexposed parts
dark red to plum. Girdle broad, fleshy, tan to dark reddish
brown, appearing smooth but with very small clusters of short,
stout, white spicules widely scattered on dorsal surface,
especially where girdle intrudes between valves; long spicules
in dorsal tufts reddish brown, shorter basal spicules of tufts
blue-green.

Valve i semilunate (Fig. 11), wider than long, concave
posteriorly, with anterior insertion plate bearing 5 U-shaped
slits; tegmentum occupying about 65% total valve length.
Valves ii-vii beaked (Fig. 12); tegmentum alate (wing-shaped),
little wider than long but constricted over much of anterior
portion, with markedly concave anterolateral margins; sutural
laminae very large, longer than wide, broad, flared antero-
laterally, separated anteriorly by wide, deep, U-shaped sinus;
lateral margins not parallel to each other or to jugum; single
slits near midpoints of margins. Valve viii broadly triangular
(Fig. 13), about twice as wide as long, rounded posteriorly,
with mucro posterior of center; tegmentum drop-shaped,
longer than wide, constricted anteriorly along jugum; sutural
laminae very wide, flared anteriorly, separated by wide, V-
shaped sinus, with straight anterior margins; 2 slits in posterior
insertion plate small, narrow, V-shaped.

Jugum smooth, narrow, with parallel sides well-
separated from lateral tegmental surface, extending anteriorly
beyond main body of tegmentum. Tegmental pustules drop-
shaped (Fig. 14), 120 um long, 80 nm wide, with single cen-
tral macresthete, 3-6 micresthetes nearly all adapical of
macresthete.

Girdle upper surface covered with dense mat of very

small (50 »m) brown spicules interrupted by clusters of stout,
white, 200-400 um long spicules (Fig. 15), clusters very sparse
on main dorsal surface, dense where girdle intrudes between
valves; 18 anterior and sutural tufts containing about 50
straight, stout, sharp-tipped spicules up to 2 mm long, brown
along shafts, blue-green at base, with extremely fine, needle-
like spicules within base; margin fringed with slender, straight
or slightly curved, sharp-tipped blue or blue-green spicules
up to 1.4 mm long; underside densely covered with slender,
sharp-tipped spicules about 80 um long, directed toward
periphery.
DISCUSSION: Pilsbry (1983) described Acanthochitona
rhodea from an alcoholic specimen 28 mm long, 15 mm wide,
that had already ‘‘lost the cuticle and hairs from its girdle,
leaving a smooth whitish surface pitted at the sutures.’ Thus,
one important identification character, the girdle spicules,
could not be described. Now all that remains of the holotype
are three disarticulated valves, ii, vii(?), and viii (Figs. 16-18).
Nevertheless, sufficient evidence remains in the drop-shaped
pustules, well-illustrated by Pilsbry (1893: pl. 12, fig. 49), to
demonstrate that A. rhodea is the species that inhabits the
Caribbean coast of Panama. Thus, Ferreira’s (1985) restric-
tion of the type locality to Portobelo was appropriate.

Characters important in separating Acanthochitona
rhodea from A. hemphilli include the drop-shaped rather than
reniform pustules, the dark red rather than greenish white
sutural laminae, and the small clusters of stout spicules widely
scattered among the mat of shorter spicules on the dorsal

Figs. 16-18. Acanthochitona rhodea (Pilsbry, 1893), holotype;
‘“‘Panama’’; ANSP 63429. Fig. 16. Valve ii. Fig. 17. Valve vii (7). Fig.
18. Valve viii.

surface of the girdle. Differences between A. rhodea and the
Pacific coast species are discussed under remarks following
the description of that species.

Ferreira’s (1985) distributional records of Acantho-
chitona rhodea are unreliable because he identifed all three
species as A. rhodea. Houbrick’s (1968) specimens from the
Caribbean coast of Costa Rica, which | examined, are A.
rhodea. Leloup’s (1941) description and illustrations of drop-
shaped tegmental pustules (his figs. 5, 6) and large spicules
scattered in widely separated groups among the smaller
spicules of the dorsal girdle surface demonstrate that his
specimens from off Colombia in 28-29 fm (51-53 m) were A.
rhodea. Scattered spicule clusters on the girdle and shapes
of valves i, vii, and viii indicate that specimens illustrated as
A. hemphilli from Curagao by Kaas (1972) are A. rhodea.
Likewise, Gotting’s (1973) illustration of scattered clusters of
girdle spicules indicates that specimens he reported as A.
rhodea from Caribbean Colombia were identified correctly.
Thus, the species is known with certainty only from the
southern Caribbean Sea, where it usually is collected in the
shallow subtidal zone.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 85

Acanthochitona ferreirai Lyons, sp. nov.
Figs. 19-24

Acanthochitona rhodea, Keen, 1958: 519, fig. 10 (pars). A. G.
Smith, 1961: 89. Thorpe /n Keen, 1971: 867, 868, fig. 14.
Bullock, 1974: 164 (pars). Ferreira, 1985: 207, 208
(pars). [non A. rhodea (Pilsbry, 1893)}.

TYPE MATERIAL: HOLOTYPE: 28.2 mm, Punta Mala, Pacific coast
of Panama, July 1969, R. C. Bullock, collector, USNM 859314.
PARATYPES: PANAMA: 13 spec., 9.4-28.2 mm, collected with
holotype, ANSP A12121 (1), CAS 064883 (1), RMNH 55985 (1), FSBC
| 32563 (2), Bullock collection (8). COSTA RICA: 2 spec., 19.6,
26.5 mm, Playa de Jaco, intertidal, 25 Apr 1975, FSBC | 32564.

TYPE LOCALITY: Punta Mala, Panama.

DISTRIBUTION: Pacific coasts of Costa Rica and Panama;
intertidal and shallow subtidal depths.

iy el
Auld 5, vot

DESCRIPTION: Largest specimen (holotype) 28.2 mm long,
17.0 mm wide including girdle; valves occupying approximate-
ly 65% of total specimen width (Fig. 19). Exposed valves
uniformly red or rose, usually with white maculations; unex-
posed parts rose pink. Girdle broad, orange-brown or dark
red, with large white patches of spicules unevenly spread
across dorsal surface; spicules of dorsal tufts green.
Valve i semilunate (Fig. 20), wider than long, concave
posteriorly, with anterior insertion plate bearing 5 slits; tegmen-
tum occupying about 65% total valve length. Valves ii-vii
beaked (Fig. 21); tegmentum alate, twice as wide as long, con-
stricted anteriorly, with anterolateral margins concave near
jugum; sutural laminae broad, flared anterolaterally, separated
anteriorly by wide, shallow sinus; lateral margins not parallel
with each other or with jugum; single slits near midpoints of
margins. Valve viii broadly triangular (Fig. 22), twice as wide
as long, rounded posteriorly, with nearly central mucro;

Figs. 19-24. Acanthochitona ferreirai Lyons, sp. nov. Fig. 19. Holotype, 28.2 mm; Punta Mala, Panama; USNM 859314. Fig. 20. Valve i ex
24.5 mm paratype; same location; FSBC | 32563. Fig. 21. Valve iv, same specimen. Fig. 22. Valve viii, same specimen. Fig. 23. Tegmental
pustules, valve iv, same specimen (field width = 280 ym). Fig. 24. Dorsal girdle spicules, same specimen (field width = 650 ym).

86 AMER. MALAC. BULL. 6(1) (1988)

tegmentum ovate, wider than long, constricted anteriorly along
jugum; sutural laminae very wide, flared anterolaterally, with
straight anterior margins, separated by very shallow, broad,
V-shaped sinus; 2 slits in posterior insertion plate small, nar-
row, V-shaped.

Jugum smooth, narrow, with parallel sides well-
separated from lateral tegmental surface, extended anterior-
ly beyond main tegmental mass. Tegmentum of all valves
covered with small (100 nm) round to slightly ovate pustules
(Fig. 23), with single subcentral macresthete, 3-4 micresthetes.

Girdle upper surface covered with dense mat of very
small (60 pm) spicules overlain by extensive patches of
slender, straight, white spicules 400-500 um long (Fig. 24),
especially evident posteriorly and where girdle intrudes be-
tween valves; 18 anterior and sutural tufts containing 50-60
straight or slightly curved, stout, sharp-tipped green spicules
up to 2.2 mm long; margin fringed with slender, sharp-tipped
spicules up to 1 mm long, arranged in alternating groups of
purple and white; underside densely covered with slender,
sharp-tipped spicules about 80-90 um long, directed toward
periphery.

DISCUSSION: Acanthochitona ferreirai is related to A.
hemphilli and, especially, to A. rhodea of the Caribbean region.
The relatively shorter, wider valves, round to subovate tegmen-
tal pustules, and dense, clearly evident patches of longer
spicules on the dorsal surface of the girdle separate A. fer-
reirai from the other two species.

This is the species reported from the eastern Pacific
as Acanthochitona rhodea by Keen (1958), A. G. Smith (1961),
Thorpe (/n Keen, 1971), Bullock (1974), and Ferreira (1985).
Together, those authors reported specimens ranging from
Mexico to Peru. | examined only specimens from Costa Rica
and Panama, so | cannot confirm that specimens from Mex-
ico and Peru are conspecific with the material described here.

ETYMOLOGY: Named for the late Antonio J. Ferreira, whose
work stimulated much interest in Caribbean and Panamic
polyplacophorans.

Acanthochitona spiculosa (Reeve, 1847)
Figs. 25-29

Chiton spiculosus Reeve, 1847: pl. 9, sp. and fig. 47.
Acanthochites spiculosus, Pilsbry, 1893: 22, pl. 13, figs. 60-62.
(non Acanthochitona spiculosa of subsequent authors).

TYPE MATERIAL: LECTOTYPE: 33.0 mm; “‘Loc. West Indies; Cum-
ing collection; Acc. 1829’’; BM (NH) 1981251/1 (herein designated).
PARALECTOTYPES: 4 spec., 21.0-28.0 mm; collected with lectotype;
BM (NH) 1981251/2-5.

DISCUSSION: All five types (Figs. 25-29) at one time were
glued to a tablet by either the dorsal or ventral surface. Three
specimens contain the dried remains of the foot and viscera,
and two have been scraped clean beneath. Within one of the
latter, a tag was glued but has been removed, leaving only
a torn remnant upon which no information remains. This
specimen [BM(NH) 1981251/1], previously labeled as the
figured syntype, is the most flattened and best preserved of
the specimens and most resembles Reeve’s figure 47 in its
proportions of length and width; it is designated herein as the
lectotype. However, if this is the specimen figured by Reeve,
considerable liberties were taken to enhance the illustration.
Reeve’s figure depicts a black, smooth, shiny surface over
all valves; each intermediate valve is drawn with a distinct
jugal separation extending obliquely from the posterior beak
to the anterolateral corners of the exposed valve surface; a
single concentric band appears near the lateral margins of
each valve. Spicules of dorsal tufts are depicted as long,
densely packed, and fully spread from each cluster, overlying
the entire girdle and extending beyond its narrow margin. The

Figs. 25-29. Acanthochitona spiculosa (Reeve, 1847), type specimens, ‘‘West Indies’’. Fig. 25. Paralectotype, 27.0 mm; BM(NH) 1981251/3.
Fig. 26. Paralectotype, 28.0 mm; BM(NH) 1981251/2. Fig. 27. Lectotype, 33.0 mm; BM(NH) 1981251/1. Figs. 28, 29. Paralectotypes, 26.0, 21.0

mm; BM(NH) 1981251/4, 5.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 87

spicules are olive with traces of blue-green.

The actual syntypes are not nearly so attractive. Ex-
pectedly, having been dried for more than 150 years, the
girdles are shrunken and hardened, and many of the spicules
are broken. However, the greatest difference between the
specimens and Reeve’s description is in the condition of the
dorsal tegmentum. The jugal tract and most of the
lateropleural areas of each valve of every specimen are
severely eroded, evidently as a result of surf abrasion (this
condition occurs frequently among Caribbean species such
as Acanthopleura granulata, Chiton squamosus, and
Ceratozona squalida which inhabit intertidal zones of surf-
swept rocks). The only remaining tegmentum occurs near
lateral margins of the valves; the intersection of original
tegmentum and eroded valve is evidently the concentric band
depicted in Reeve’s figure. On some specimens, the jugal area
is marked by an eroded dark band set apart by lighter areas
at each side, but the only actual remnants of jugum were found
beneath the overhang of the posterior edge of the preceding
valve on valve ii of the smallest curled specimen and on valve
viii of the next larger curled specimen. The jugum is black,
nearly smooth but microscopically pitted. No incisions are evi-
dent on the jugum of any syntype. The densely arranged
pustules near lateral margins of intermediate valves are so
coated with grime that their form is difficult to discern, but,
where apparent, they vary from ovate to drop-shaped.

Four types of spicules occur on the girdle. Those of
the 18 dorsal tufts, although frequently broken, are most evi-
dent, being long, round, sharp-pointed, and densely packed;
individual lengths vary considerably, as do corresponding
thicknesses; their color is now light golden brown. Aside
from the tufts, the dorsal surface of the girdle is covered with
short, blunt-tipped, club-shaped brown spicules. Fairly short,
slender, vitreous, sharp-pointed spicules form a fringe at the
outer margin of the girdle. On the underside, densely packed,
very short, vitreous spicules barely break the girdle surface.
Particles of quartz sand occur among debris trapped in gir-
dle spicules of the types.

The taxonomic history of Acanthochitona spiculosa has
been greatly confused. Much of that confusion can be traced
to W. H. Dall and E. A. Smith. Dall (1889a) identified as
A. spiculosa specimens hereafter shown to be A. pygmaea
(Pilsbry, 1893). In the following year, E. A. Smith (1890) com-
bined A. spiculosa with A. astrigera (Reeve, 1847). Pilsbry
(1893) included A. spiculosa among species in the West
Indian fauna, but he only attempted to reproduce Reeve’s
description verbally and visually and did not report any addi-
tional material. A full synonymy of correct and incorrect ap-
plications of the name A. spiculosa, and of its confusion with
A. astrigera, A. pygmaea, and other taxa, comprises nearly
five manuscript pages. Because most references cited are
lists or repetitions of relatively few uncritical but far-reaching
decisions, only the more important are discussed in the follow-
ing species accounts.

Valve morphology and other characters of the syntypes
indicate that Acanthochitona spiculosa is related to the group
containing the Caribbean A. astrigera, the eastern Pacific
A. hirudiniformis (Sowerby, 1832), and the Hawaiian A. viridis

(Pease, 1872). However, the syntypes are so worn that they
cannot be related with certainty to any of those species. The
valves are somewhat wider and the dorsal tuft spicules are
shorter, more coarse, and less numerous than are those of
both A. astrigera and another Caribbean species described
hereafter. The valves and spicules of A. spiculosa seem to most
resemble those of A. hirudiniformis; if they prove to be con-
specific, the latter name will have priority. No other specimens
of Caribbean, Brazilian, or East Pacific Acanthochitona resem-
ble the syntypes of A. spiculosa. Until the syntypes can be
related with certainty to specimens of known locality, A.
spiculosa should be considered a species inquirenda.

Acanthochitona astrigera (Reeve, 1847)
Figs. 30-41

Chiton astriger Reeve, 1847: pl. 18, sp. and fig. 109.

Acanthochiton astriger, Dall, 1889a: 174, 175.

(?) Chiton (Acanthochiton) astriger, E. A. Smith, 1890: 496, 497.

Acanthochites spiculosus var. astriger, Pilsbry, 1893: 22, 23,
pl. 13, figs. 55-57.

Acanthochitona spiculosa, Kaas, 1972: 46-49 (pars) non A.
spiculosa (Reeve, 1847).

Acanthochitona astriger, Watters, 1981: 173 (pars, non pl. 2d,
pl. 4h).

Acanthochitona astrigera, Lyons, 1983: 91. Ferreira, 1985:
205-207 (pars).

TYPE MATERIAL: LECTOTYPE: 19.0 mm, Barbados, BM(NH)
19809/4 (herein designated). PARALECTOTYPES: 3 spec., 19.5-22.0
mm, collected with lectotype, BM(NH) 19809/1-3.

OTHER MATERIAL EXAMINED: BAHAMAS: 30 spec., 11.5-
19.0 mm, Eight Mile Rock, Grand Bahama, intertidal, 21-23 May 1981,
FSBC | 32527. —3 spec., 9.4-12.7 mm, Bartlett Hill, Eight Mile Rock,
Grand Bahama, intertidal, 29 Aug 1984, FSBC | 32038. —28 spec.,
9.9-21.5 mm, Hunters, Grand Bahama, intertidal, 29 Aug 1984, FSBC
| 32037. —5 spec., Silver Cove Canal, Freeport, Grand Bahama, 0.5-
1.5 m, 28 Aug 1984, FSBC | 32036. —1 spec., curled, Grand Bahama,
RMNH K3730. —1 spec., Chub Cay, intertidal, M. Williams collec-
tion. DOMINICAN REPUBLIC: 3 valves, Playa Embassy, 16 km east
of Boca Chica, beach drift, Bullock collection. CAYMAN ISLANDS:
2 spec., 14.0, 15.0 mm, Jackson’s Point, Grand Cayman, 0-0.5 m, 9
June 1973, RMNH. ST. MAARTEN: 1 spec., 16.9 mm, W. Long Beach,
RMNH K4952. BARBADOS: 1 spec., 23.5 mm, Archers Bay, St. Lucy,
5 Sept 1970, Bullock collection. BONAIRE: 7 spec., 11.7-25.2 mm,
Kralendijk, intertidal, 9 Oct 1986, FSBC | 32528. CURACAO: 2 spec.,
13.5, 18.7 mm, Port Marie, 16-18 Apr 1966, Bullock collection.

TYPE LOCALITY: Barbados (original designation).

DISTRIBUTION: Grand Bahama Island to Grand Cayman
Island, Barbados, Bonaire, and Curagao; intertidal or very
shallow depths.

DESCRIPTION OF TYPES: All four types (Figs. 30-33)
previously glued to tablet, later removed; 3 glued by ventral
surface, 1 by dorsal surface. Foot and viscera remaining in
all specimens.

Overall shape elongate, relatively slender, with dimen-
sions 22 x 8 mm, 22 x 10 mm, 19.5 x 8 mm, 19 x 7.5 mm.

88 AMER. MALAC. BULL. 6(1) (1988)

Figs. 30-33. Acanthochitona astrigera (Reeve, 1847), type specimens;
Barbados. Fig. 30. Paralectotype, 22.0 mm; BM(NH) 19809/1. Fig.
31. Lectotype, 19.0 mm; BM(NH) 19809/4. Figs. 32, 33. Paralectotypes,
19.5, 22.0 mm; BM(NH) 19809/3, 2.

Three of four specimens encrusted to varying degrees by cor-
alline algae, although some valves have been cleaned. Valves
of smallest specimen in excellent condition.

Girdle brown, encroaching over anterolateral areas of
valves so that intermediate valves are shield-shaped. Valve
color varying from brown with white maculations laterally to
dark blue-green, approaching black; where tegmentum
damaged, underlying shell blue-green. Tegmentum covered
with small pustules, drop-shaped near jugum, more ovate near
center. Pustules of valve i small, ovate near apex, larger, drop-
shaped near margins. Jugum of intermediate valves slender,
with nearly smooth surface rendered finely striate by linear
arrangement of fine pits near margins, pits exposed across
entire jugal width near beaks; subsurface striations visible
through smooth jugum surface in remaining areas. Valve viii
with drop-shaped to ovate pustules as on other valves; mucro
relatively low, posterior of center; jugum smooth, but with
longitudinal striae visible beneath transparent surface.

Spicules of anterior and sutural girdle tufts extremely
dense, white, straight, very slender; numerous very small

spicules on dorsum of girdle; spicules at girdle margin stout,
long, approximately 3 length of those in tufts, overlying
shorter, sharp-tipped spicules, both types glassy, white; under-
side of girdle with very fine, short spicules protruding through.
Fragments of foraminifera and carbonate particles trapped
among girdle spicules.

SUPPLEMENTAL DESCRIPTION: Largest specimen (FSBC
| 32528) 25.2 mm long, 15.0 mm wide; valves occupying about
30% total specimen width (Fig. 34). Valves dark blue-green
to black, usually with white, stripe-like maculations on valves
ii and v, less commonly on other valves. Girdle blue-green,
brown, or black, virtually obscured by expanded tufts of long,
slender spicules.

Valve | semilunate (Fig. 35), wider than long, posterior
margin straight, with anterior insertion plate bearing 5 slits;
tegmentum occupying approximately 70% of valve length. In-
termediate valves ii-vii beaked posteriorly, with smooth jugum
widening anteriorly (Figs. 36, 37); tegmentum pentagonal, as
long as wide in all but smallest specimens, with straight to
slightly convex anterolateral margins; sutural laminae large,
broad, curving anteromedially from posterolateral corners of
tegmentum, with broadly to acutely rounded tips separated
by broad sinus; single narrow slits along lateral margins. Valve
viii tegmentum trigonal (Figs. 38-40), widest anteriorly, with
anterior margin straight at broad sinus; mucro elevated,
posterior of center; sutural laminae very well-developed,
broad, slightly or markedly sinuous along margins; two slits
in posterior insertion plate distinct, varying in width and depth.
Valve viii often misshapen, asymmetrical or missing features
(Figs. 39, 40). Pustules of tegmentum ovate to drop-shaped
(Fig. 41), shallowly cupped, 80-90 »m long, constricted
adapically, with single, large, macresthete, 1-3 micresthetes
at juncture of apex and tegmental plain.

Girdle upper surface dominated by 18 anterior and
sutural tufts each comprised of more than 100 white to light
amber, long (to 4 mm), slightly curved, slender, sharp-tipped
spicules; background spicules of dorsal surface short (100
pm), straight, sharp-tipped, blue, brown, or black; marginal
spicules white, approximately 500 pm long, straight, slender,
sharp-tipped; underside covered with fine (80 um), sharp-
tipped spicules directed toward periphery.

DISCUSSION: The dark blue-green valves, densely packed,
long, slender, sharp-tipped spicules of anterior and sutural
girdle tufts, and white maculations on the blue-green tegmen-
tum, often only on valves ii and v, leave no doubt that
specimens reported here as Acanthochitona astrigera are con-
specific with those described by Reeve. Bullock’s 23.5 mm
specimen from Barbados is identical in all respects with
Reeve’s syntypes. Reeve’s smallest specimen, illustrated in
Fig. 31, is designated herein as lectotype.

E. A. Smith (1890) initiated the confusion between
Acanthochitona astrigera and A. spiculosa with the statement:
“‘TReeve’s] figure (47) of the detail of sculpture of C. spiculosa,
Reeve, which | believe to be the same species [as C.
astrigera], gives quite as good an idea of the ornamentation
{of astrigera] as [Reeve’s] figure 109’ Pilsbry’s (1893)
diagnostic comments indicate that he correctly recognized

LYONS: CARIBBEAN ACANTHOCHITONIDAE 89

Figs. 34-41. Acanthochitona astrigera (Reeve, 1847). Fig. 34. Whole specimen, 20.2 mm; Eight Mile Rock, Grand Bahama; FSBC | 32527.
Fig. 35. Valve i ex 15.0 mm specimen; same lot. Fig. 36. Valve iv, same specimen. Fig. 37. Valve v ex 18.1 mm specimen; same lot. Fig.
38. Valve viii ex 12.2 mm specimen, same lot. Fig. 39. Valve viii, same specimen as 35. Fig. 40. Valve viii, same specimen as 37. Fig. 41.
Tegmental pustules, valve iv, same specimen as 38 (field width = 280 pm).

A. astrigera, but he cited Smith as authority in designating
astrigera a variety of A. spiculosa despite the fact that Smith
chose to use astrigera, not spiculosa, for his material. Pilsbry
cited no material of A. spiculosa s.s. Thereafter, A. astrigera
was reported by many authors under the name A. spiculosa,
as were many specimens of A. pygmaea and other taxa. Kaas
(1972) reported as A. spiculosa specimens of A. astrigera from
Grand Bahama Island, but he also reported some specimens
of A. pygmaea as A. spiculosa (see remarks for that species).
Watters (1981) correctly noted that A. astrigera and A.
spiculosa were distinct, but he included an undescribed
species within his concept of A. astrigera, and he supported
Dall’s misconception that A. spiculosa represented the
species otherwise known as A. pygmaea (Pilsbry, 1893). Fer-
reira (1985) followed Watters’ concepts of both A. astrigera
and A. spiculosa.

Thus, published records of Acanthochitona spiculosa
and A. astrigera, its supposed synonym, actually include
A. astrigera, A. pygmaea, and, according to material | have
examined, some specimens of A. andersoni Watters, 1981,
and a new species described hereafter. Specimens illustrated
as A. astrigera by Watters (1981) from La Parguera, Puerto
Rico, and Water Id., Virgin Islands, are the new species.
Likewise, Ferreira’s (1985) record of A. astrigera from Belize
was based upon an IRCZM specimen of the new species. The
literature can be corrected only when previously reported
specimens, including E. A. Smith’s Fernando Noronha record,
have been re-examined.

Dall (1889a) listed Acanthochitona astrigera from Dry
Tortugas and the Florida Keys, but | have seen no specimens
from Florida. At Grand Bahama Island, A. astrigera lives prin-
cipally among brown algae in the intertidal zone of high wave

90 AMER. MALAC. BULL. 6(1) (1988)

energy, rocky shores, a habitat absent from the Florida Keys.

The relationship of Acanthochitona astrigera to other
New World Acanthochitona is discussed under remarks for
the new species.

Acanthochitona lineata Lyons, sp. nov.
Figs. 42-51

Acanthochitona astriger, Watters, 1981 (pars, pl. 2d, pl. 4h).
Acanthochitona astrigera, Ferreira, 1985: 206-208 (pars,
Belize). [non A. astrigera (Reeve, 1847)].

TYPE MATERIAL: HOLOTYPE: 19.5 mm x 10.5 mm, Silver Cove
Canal, Freeport, Grand Bahama Island, 0.5-1.5 m, 28 Aug 1984, W.
G. Lyons, collector, USNM 859315. PARATYPES: BAHAMAS: 1 spec.,
10.8 mm, Bartlett Hill, Eight Mile Rock, Grand Bahama, 0-0.5 m, 29
Aug 1984, FSBC | 32434. —34 spec., 5.6-22.5 mm, same locality and
date as holotype, ANSP A12122 (2), RMNH 55986 (2), FSBC | 32433
(29). —2 spec., 22.6, 33.0 mm, McLeanstown, east end Grand
Bahama, 1 m, 27 Aug 1984, FSBC | 32432. PUERTO RICO: 4 spec.,
7.0-11.1 mm, Magueyes Id., La Parguera, 1967, Bullock collection. —16
spec., 8.5-13.9 mm, Magueyes Id., Bullock collection (12), FSBC |
32565 (4). —1 spec., 21.4 mm, Media Luna Reef, La Parguera, 0-2 m,
15 Aug 1985, FSBC | 32435. —1 spec., 21.8 mm, Cayo Enrique, La
Parguera, 0-1 m, 19 Aug 1985, FSBC | 32436. VIRGIN ISLANDS: 1
spec., 9.7 mm, Water lId., July 1959, DMNH 95381. BELIZE: 1 spec.,
20.0 mm, Carrie Bow Cay, 0-1 m, 23 Mar 1981, IRCZM 61:052.

TYPE LOCALITY: Silver Cove Canal, Freeport, Grand
Bahama Island.

DISTRIBUTION: Grand Bahama Island to Puerto Rico, the
Virgin Islands, and Belize, 0.5-2.0 m.

DESCRIPTION: Largest specimen 33.0 mm long, 18.0 mm
wide including girdle; valves occupying approximately 30%
of total specimen width (Figs. 42-44). Valve i with 6-8
olivaceous, equally spaced concentric lines, expressed on
valves ii-vii as 3-7 transverse stripes (chevrons) extending
posterolaterally from jugum; stripes varying in strength and
number among individual specimens, usually strongest, most
numerous, on valves i-iii; valves iii-v occasionally dark green,
brown, or black, obscuring stripes; valve viii mostly white, with
single large spots on lateral areas. Girdle entirely white or buff,
sometimes mottled with brown or blue-green bands, occa-
sionally with large brown or black spot near middle.

Valve i semilunate (Fig. 45), wider than long, posterior
margin straight, with anterior insertion plate bearing 5 slits;
tegmentum occupying 80-90% of valve length. Valves ii-vii
strongly produced posteriorly; tegmentum pentagonal, with
straight to slightly convex anterolateral margins (Fig. 46);
sutural laminae well-developed, curving anteromedially from
posterior corners of tegmentum, with subacute anterior tips
separated by relatively narrow sinus; single slits on lateral
margins. Valve viii tegmentum ovate (Fig. 47), widest latero-
mesially; mucro elevated, slightly posterior of center; sutural
laminae well-developed, broadly subquadrate; two slits in
posterior insertion plate very large. Proportions of small
specimens may differ from those of larger individuals (Figs.
48-50).

AA

Figs. 42-44. Acanthochitona lineata Lyons, sp. nov. Fig. 42. Holotype, 19.5 mm; Freeport, Grand Bahama; USNM 859315. Fig. 43. Paratype,
13.1 mm; La Parguera, Puerto Rico; FSBC | 32565. Fig. 44. Paratype, 33.0 mm; McLeanstown, Grand Bahama; FSBC | 32432.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 2

Jugum smooth, relatively narrow on valves ii-vii, wide
anteriorly on valve viii. Tegmentum of all valves covered evenly
with small (50 xm), round to slightly ovate, shallowly cupped
pustules (Fig. 51) with single, central macresthete, 1-2
micresthetes near apex.

Girdle upper surface appearing smooth, actually
covered with extremely fine (20-30 xm) spicules; 18 anterior
and sutural dorsal tufts comprised of more than 100 white,
occasionally amber, long (to 3.5 mm), straight, very slender,
sharp-tipped spicules; marginal spicules white, approximately

“ a,

Figs. 45-51. Acanthochitona lineata Lyons, sp. nov. Fig. 45. Valve i ex 16.5 mm paratype; Freeport, Grand Bahama; FSBC | 32433. Fig. 46.
Valve iv, same specimen. Fig. 47. Valve viii, same specimen. Fig. 48. Valve i ex 12.0 mm paratype; La Parguera, Puerto Rico; FSBC | 32565.
Fig. 49. Valve iv, same specimen. Fig. 50. Valve viii, same specimen. Fig. 51. Tegmental pustules, valve iv, same specimen (field width = 280 pm).

800 um iong, straight, slender, sharp-tipped; underside
covered with fine (80 um), sharp-tipped spicules directed
toward periphery.

DISCUSSION: Acanthochitona lineata is related closely to A.
astrigera of the Caribbean Sea and to A. hirudiniformis (Sower-
by, 1832) (Figs. 52-56) of the tropical eastern Pacific Ocean;
valves of all three species are quite similar. However, the
tegmentum of valve viii of A. astrigera is widest anteriorly,
whereas those of A. lineata and A. hirudiniformis are widest

92 AMER. MALAC. BULL. 6(1) (1988)

Figs. 52-56. Acanthochitona hirudiniformis (Sowerby, 1832). Fig. 52. Whole specimen, 23.0 mm; Playa de Jaco, Costa Rica; FSBC | 32566.
Fig. 53. Valve i ex 14.5 mm specimen, same lot. Fig. 54. Valve iv, same specimen. Fig. 55. Valve viii, same specimen. Fig. 56. Tegmental

pustules, valve iv, same specimen (field width = 265 um).

mesially. In addition, tegmental pustules of A. astrigera are
drop-shaped, whereas pustules of the other two species are
round, those of A. hirudiniformis being approximately 50%
larger than those of A. lineata on specimens of similar size.
Other differences which separate A. hirudiniformis from A.
lineata include the longer spicules of the girdle mat, which
give a rough rather than smooth appearance to the dorsal
surface, the short green greater than long white spicules of
the anterior and sutural tufts, and the diffuse rather than clear-
ly demarked color pattern on the tegmentum. Like A. astrigera,
A. hirudiniformis lives intertidally on high energy rocky shores,
whereas A. lineata usually occupies shallow, subtidal, relative-
ly more placid areas such as reef flats.

Ferreira (1985) identified the IRCZM specimen of Acan-
thochitona lineata from Carrie Bow Cay, Belize, as A. astrigera.
The specimen from Water Id., Virgin Islands (DMNH 95381,
not 45381) illustrated as A. astrigera by Watters (1981: 176,
pl. 4h) also is A. lineata.

Specimens of Acanthochitona lineata | examined
seldom exceeded 22 mm length. The 33 mm specimen (FSBC
| 32432) was collected among many large (30-50 mm) A.
hemphilli at the base of a colony of finger coral, Porites
astreoides.

ETYMOLOGY: From Latin, ‘‘linea’’, to denote the lines or
stripes on the tegmentum.

Acanthochitona worsfoldi Lyons, sp. nov.
Figs. 57-65

(?) Choneplax cf. lata, Ferreira, 1985: 208-213 (pars, figs. 16,
17). [non Choneplax lata (Guilding, 1829)].

TYPE MATERIAL: HOLOTYPE: Length 14.8 mm, width 6.7 mm,
Silver Cove Canal, Freeport, Grand Bahama Island, 0.5-1.5 m, 28 Aug
1984, W. G. Lyons, collector, USNM 859318. PARATYPES: BAHAMAS:
6 spec., 7.7-17.2 mm, same locality and date as holotype, ANSP A12123
(1), FSBC | 32545 (5). —2 spec., 9.0, 13.6 mm, Tamarind Beach Reef,
Grand Bahama, 18 m, 28 Aug 1984, RMNH 55987 (1), FSBC | 32544
(1). —2 spec., 7.0, 10.1 mm, Tamarind Beach Reef, 39 m, Sept 1983,
FSBC | 32543. —1 spec., 13.5 mm, Gold Rock, Grand Bahama,
24.4 m, 1980, FSBC | 32541. —5 spec., 8.5-12.0 mm, Gold Rock,
24.4 m, Aug 1983, FSBC | 32542. —1 spec., 10.0 mm, 2 km off Bell
Channel, Lucaya, Grand Bahama, 18.3-19.8 m, 6 Apr 1974, FSBC |
32539. —1 spec., 12.0 mm, 2 km off Bell Channel, Lucaya, 45.7 m,
10 July 1974, FSBC | 32540.

OTHER MATERIAL EXAMINED: 8 valves, Gold Rock, bottom
sediments, 24.4 m, May-July 1981, FSBC | 32534. —53 valves, Grand
Bahama, bottom sediments, May 1981, R. Quigley collection.

TYPE LOCALITY: Silver Cove Canal, Freeport, Grand
Bahama Island.

DISTRIBUTION: Grand Bahama Island, 0.5-45.7 m, ?
Barbados.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 93

WA am

NY)

a

agit’

\

fe
oF

Figs. 57-65. Acanthochitona worsfoldi Lyons, sp. nov. Fig. 57. Holotype, 14.8 mm; Freeport, Grand Bahama; USNM 859318. Fig. 58. Paratype,
17.2 mm; same location; FSBC | 32545. Fig. 59. Valve i ex 12.0 mm paratype; Gold Rock, Grand Bahama; FSBC | 32542. Fig. 60. Valve iv,
same specimen. Fig. 61. Valve viii, same specimen. Fig. 62. Valve i ex 80 mm paratype; Freeport, Grand Bahama; FSBC | 32545. Fig. 63.
Valve iv, same specimen. Fig. 64. Valve viii, same specimen. Fig. 65. Tegmental pustules, valve iv, same specimen (field width = 250 um).

DESCRIPTION: Largest specimen 17.2 mm long, 7.4 mm wide
including girdle; valves occupying about 40% of total
specimen width (Figs. 57, 58). Valves highly arched, orange,
rust, or bright red, with scattered white maculations on
tegmentum. Girdle buff, usually crossed with reddish brown
bars which continue onto spicular fringe at margin.

Valve i semilunate (Fig. 59), wider than long, posterior
margin straight, with anterior insertion plate bearing 5 shallow
slits; tegmentum occupying approximately 80% of valve
length. Valves ii-vii beaked posteriorly (Fig. 60); tegmentum
pentagonal to subcircular, rounded anteriorly, about as long
as wide, with convex anterolateral margins; sutural laminae

small, curving anteromedially from posterior corners of
tegmentum; subacute anterior tips separated by wide, shallow
sinus; single, small, narrow slits along lateral margins. Valve
viii tegmentum subovate (Fig. 61), widest lateromesially, with
straight anterior margin; mucro elevated, posterior of center;
sutural laminae subrectangular, as wide or wider than long;
two slits in posterior insertion plate small, V-shaped. Propor-
tions of small specimens may differ from those of larger in-
dividuals (Figs. 62-64).

Jugum smooth, narrow at beaks, expanded anterior-
ly. Tegmentum of all valves covered evenly with small
(50-60 um), flattened subspatulate pustules (Fig. 65) with

94 AMER. MALAC. BULL. 6(1) (1988)

single subapical macresthete, 1-2 micresthetes at apex.

Girdle upper surface appearing smooth, actually
covered with fine (50 um) spicules; 18 anterior and sutural
dorsal tufts comprised of 10-15 long (1.5 mm), slender, slight-
ly curved, sharp-tipped, reddish brown or white spicules;
margin densely fringed with long (1.0-1.2 mm), slender, slightly
curved, sharp-tipped spicules similar to those in dorsal tufts;
underside covered with fine (80 nm), narrow, straight, sharp-
tipped spicules directed toward periphery.

DISCUSSION: Acanthochitona worsfoldi occurs in two color
forms. The typical form, exemplified by the holotype (Fig. 57),
has rusty orange valves, girdle, and spicules of the dorsal tufts
and marginal fringe. Another form, represented by single
specimens from 0.5-1.5 m and 38.0 m depths, has bright red
valves, a light buff girdle, and only clear, vitreous spicules in
the dorsal tufts and marginal fringe (Fig. 58). The two forms
are identical morphologically.

Acanthochitona worsfoldi is distinguished from other
species by its combination of large, subcircular tegmentum,
small sutural laminae, few spicules in dorsal tufts, and dense
marginal fringe of large spicules. Valve morphology suggests
relationship to the species complex containing A. astrigera
and A. lineata, but tegmental pustules and girdle spicules of
those species differ considerably from those of A. worsfoldi.

The bathymetric range of Acanthochitona worsfoldi
generally is greater than that of other Caribbean Acantho-
chitona species; eight of the nine lots examined were collected
by divers using SCUBA. Ferreira (1985) diagnosed and il-
lustrated specimens from Barbados which he tentatively
assigned to Choneplax lata. | was unable to obtain that
material for examination, but Ferreira’s account suggests that
the specimens are A. worsfoldi; if so, the range of A. wors-
foldi would be extended considerably.

ETYMOLOGY: Named for Jack N. Worsfold, teacher and
naturalist extraordinaire of Grand Bahama Island, whose col-
lecting efforts contributed invaluably to many studies of marine
invertebrates, including the present work.

Acanthochitona pygmaea (Pilsbry, 1893)
Figs. 66-72

Acanthochiton spiculosus, Dall, 1889a: 174, 175 (pars). [non
A. spiculosa (Reeve, 1847)].

Acanthochites pygmaeus Pilsbry, 1893: 23, pl. 13, figs. 58, 59.

Acanthochiton pygmaeus, Leloup, 1941: 37, figs. 2, 3,
pl. 1, fig. 1 (? pars).

Acanthochiton spiculosa, Kaas, 1972: 46-49, figs. 74-81 (pars).
Watters, 1981: 173-176, pl. 2a-c, pl. 4f, g. Ferreira,
1985: 214 (pars).

Acanthochitona pygmaea, Kaas, 1972: 49, 50, figs. 82-89
(? pars).

TYPE MATERIAL: PARALECTOTYPE: approximately 8.0 mm, par-
tially disarticulated; Cedar Keys, Florida; ANSP 35782.

OTHER MATERIAL EXAMINED: FLORIDA: 27 spec., 6.4-
14.1 m, St. Andrews Bay, Panama City, 1.3-2.0 m, Jan 1982, R.
Granada collection (23), FSBC | 32474(4). —2 valves, Florida Mid-

dle Ground, 28°935’N, 84°18’W; bottom sediments, 25.6-38.1 m, 7 Mar
1976, FSBC | 32524. —9 spec., 6.4-13.1 mm, Cedar Keys, CAS 063316.
—1 spec., off Crystal River, 1.8 m, 25 Mar 1968, FSBC | 6524. —2
spec., 11.0, 11.9 mm, Anclote Key, 11 Feb 1982, FSBC | 32063. —2
spec., 4.5, 7.4 mm, 6 km west of Anclote Key, 29 Sept 1982, FSBC
| 32476. —28 spec., 2.0-9.0 mm, south end Anclote Key, 3-4 m, 1 Feb
1982, FSBC | 32475. —7 spec., 2.0-9.0 mm, south end Anclote Key,
3.5 m, 22 Sept 1982, FSBC | 32473. —5 spec., all curled, Gulfport,
RMNH K3731. —6 spec., 12.9-15.5 mm, Tampa Bay, 0.5 m, 9 July 1978,
FSBC | 32052. —16 spec., 4.5-16.6 mm, Sarasota Bay, 4 m, CAS
063318. —1 spec., curled, Charlotte Harbor, 2m, FSBC | 8457. —9
spec., 5.0-11.1 mm, Punta Rassa, 4 m, CAS 063320. —32 lots, 544
spec., Hourglass Stations B, C, J, K (18-37 m) off St. Petersburg and
Sanibel Id., eastern Gulf of Mexico, 1965-67. —8 spec., 4.0-9.8 mm,
Key West, CAS 063321. —1 spec., 18.0 mm, No Name Key, CAS
063319. —3 spec., 6.0-12.0 mm, West Summerland Key, 0-1 m, 27
Sept 1981, FSBC | 32062. —3 spec., 9.1-13.0 mm, West Summerland
Key, 1976, Bullock collection. —13 spec., 5.2-15.0 mm, West Sum-
merland Key, 1978, Bullock collection. —1 spec., 10.0 mm, Sister
Creek, Vaca Key, 0-1 m, 4 Oct 1979, FSBC | 32058. —7 spec., 4.5-
9.4 mm, Sister Creek, 0.5-1.5 m, 5 Aug 1980, FSBC | 32060. —5 spec.,
8.7-14.0 mm, north side Vaca Key, 0-1 m, 30 Sept 1979, FSBC | 32053.
—4 spec., 10.7-13.8 mm, 1 spec., disarticulated, Bonefish Key, RMNH
K2852. — 5 spec., 10.0-12.0 mm, Bonefish Key, CAS 063322. —1
spec., curled, Burnt Point, Crawl Key, 2.5 m, July 1982, FSBC | 32471.
—1 spec., curled, Burnt Point, 4 Aug 1982, FSBC | 32472. —1 spec.,
10.2 mm, northeast end Grassy Key, 0.5 m, 1 Oct 1979, FSBC | 32054.
—1 spec., 11.6 mm, north side Grassy Key, 0-1 m, 1 Oct 1979, FSBC
| 32055. —12 spec., 6.2-10.7 mm, north side Grassy Key, 0.5-1.0 m,
5 Aug 1980, FSBC | 32061. —54 spec., 6.5-17.0 mm, Grassy Key
Quarry, 0-2 m, Feb 1975-Aug 1980, 4 lots: FSBC | 32051, 32056, 32057,
32059. —19 spec., 6.2-17.2 mm, Duck Key, 23 Aug 1978, Bullock col-
lection. —1 spec., 12.6 mm, Lower Matecumbe Key, CAS 063317. —1
spec., curled, off Hutchinson Id., 11.2 m, 17 Sept 1973, FSBC | 32523.
—1 spec., 11.5 mm, Bethel Shoal, 9-15 m, 27 June 1978, IRCZM 61:014.
BERMUDA: 2 spec., 12.4, 15.2 mm, Baileys Bay, July 1969, FSBC
| 32522. BAHAMAS: 2 spec., 11.5, 12.0 mm, Deadmans Reef, Grand
Bahama, 0.5-1.5 m, 25 May 1981, FSBC | 32469. —3 spec., 4.2-6.5
mm, West Hawksbill Creek, Grand Bahama, 28 June 1981, FSBC
| 32470. —12 spec., 6.7-10.7 mm, Tamarind Beach Reef, Grand
Bahama, 18 m, 28 Aug 1984, FSBC | 32064. —2 valves, Gold Rock,
Grand Bahama, bottom sediments, 24.4 m, May-July 1981, FSBC |
32525. —9 valves, Grand Bahama, bottom sediments, May 1981, R.
Quigley collection. —1 spec., 9.5 mm, McLeanstown, east end Grand
Bahama, 1-2 m, 24 May 1981, FSBC | 32468. —1 spec., 14.0 mm,
Green Turtle Cay, Abaco, 0.5 m, 3-9 May 1978, FSBC | 32466. —1
spec., 4.5 mm, Turtle Rocks near Bimini, 5.5 m, ANSP 325864. TURKS
AND CAICOS ISLANDS: 1 spec., 21.0 mm, Providenciales, M.
Williams collection. PUERTO RICO: 22 spec., 6.0-17.0 mm, Cayo Enri-
que, La Parguera, Apr 1966, Bullock collection. —8 spec., 8.5-17.7
mm, Cayo Enrique, 0.5-1.0 m, 15 Aug 1985, FSBC | 32066. —6 spec.,
4.7-16.1 mm, Cayo Enrique, 0-1 m, 19 Aug 1985, FSBC | 32069. —1
spec., 10.0 mm, Cayo Enrique, May 1985, FSBC | 32071. —2 spec.,
15.2, 179 mm, 3 km east of La Parguera, 1 m, 17 Aug 1985, FSBC
| 32067. —1 spec., 11.8 mm, Media Luna Reef, La Parguera, May 1985,
FSBC | 32070. —51 spec., 8.2-21.2 mm, Media Luna Reef, 0-2 m,
15-19 Aug 1985, FSBC | 32065. —3 spec., 8.4-20.2 mm, Isla Turramote,
La Parguera, 0-2 m, 19 Aug 1985, FSBC | 32068. VIRGIN ISLANDS:
1 spec., disarticulated, St. Thomas, RMNH K4686. SABA BANK : 2
spec., 7.5, 9.0 mm, 17°12’N, 63°38’W, 26 m, 8 June 1972, RMNH. MEX-
ICO: 2 spec., 5.0, 6.0 mm, 7 valves, Yucum Balam, 15 km north of
Ciudad Campeche, TUDG collection. —4 valves, beach 19 km
southwest of Champton, Campeche, TUDG collection. —4 valves,
Isla Arenas, 80 km north of Ciudad Campeche, TUDG collection. —2

LYONS: CARIBBEAN ACANTHOCHITONIDAE 8)

valves, Punta Palmar Lighthouse, Yucatan, TUDG collection. —7
valves, Isla Cerritos, 5 km west of San Felipe, Yucatan, TUDG col-
lection. —1 spec., 13.0 mm, Isla Mujeres, Quintana Roo, 0-1 m, 29
Sept 1985, FSBC | 32072.

TYPE LOCALITY: Key West, Florida (by subsequent designa-
tion, Watters, 1981).

DISTRIBUTION: Bermuda, both coasts of Florida, Campeche
to Quintana Roo, Mexico; Bahama Islands to Puerto Rico,
Virgin Islands, and Saba Bank; intertidal to about 40 m.

DESCRIPTION: Largest specimen 21.2 mm long, 12.0 mm
wide including girdle; valves occupying 40-45% total
specimen width (Fig. 66). Valves green, orange, often white
variegated with green or brown. Girdle buff or tan, usually
with green, blue-green or black bars, sometimes with orange
spots; dorsal spicular tufts green, blue-green or white; spicules
of marginal fringe white, usually in combination with blue or
magenta.

Valve i semilunate (Fig. 67), wider than long, broadly

V-shaped or concave posteriorly, with anterior insertion plate
bearing 5 slits; tegmentum occupying about 90% of valve
length. Valves ii-vii beaked posteriorly (Figs. 68, 69); tegmen-
tum ovate, about 1.6 times as wide as long, with convex
anterolateral margins; sutural laminae with rounded to
subacute anterior tips separated by broad sinus; single slits
along lateral margins. Valve viii trigonal (Fig. 70), widest at
anterolateral tips, rounded posteriorly; tegmentum ovate,
slightly wider than long; mucro prominent, subcentral; sutural
laminae flared anterolaterally, with straight or concave anterior
margins; 2 narrow slits in posterior insertion plate.

Jugum expanded anteriorly, with distinct longitudinal
incisions usually over entire length, sometimes rubbed smooth
anteriorly, lateral margins irregularly merging with pustules
of tegmentum. Tegmental pustules shallowly cupped, ovate
to drop-shaped (Fig. 71), about 120 1m long, 70 um wide, with
central macresthete, 3-6 micresthetes.

Girdle upper surface covered densely with slender,
vitreous, sharp-tipped spicules about 100-150 um long; 18

Figs. 66-72. Acanthochitona pygmaea (Pilsbry, 1893). Fig. 66. Whole specimen, 12.2 mm; Grassy Key, Monroe County, Florida; FSBC | 32056.
Fig. 67. Valve i ex 13.2 mm specimen; Tampa Bay, Florida; FSBC | 32052. Fig. 68. Valve iv, same specimen. Fig. 69. Valve v, same specimen.
Fig. 70. Valve viii, same specimen. Fig. 71. Tegmental pustules, valve iv, same specimen (field width = 235 um). Fig. 72. Paralectotype,

8.0 mm; Cedar Keys, Florida; ANSP 35782.

96 AMER. MALAC. BULL. 6(1) (1988)

anterior and sutural tufts comprised of 100 or more very
slender, straight, sharp-tipped spicules to 2.2 mm long; margin
fringed with straight, slender, vitreous, sharp-tipped spicules
to 700 um long; underside covered with short (80 »m), sharp,
vitreous spicules directed toward periphery.

DISCUSSION: Pilsbry (1893) described Acanthochitona
pygmaea based upon specimens from Cedar Keys and Key
West, Florida; his illustrations (pl. 13, figs. 58, 59) were of a
single intermediate valve with strongly incised jugum and an
enlarged view of tegmental pustules. Watters (1981) published
a photograph of an intact 9 mm specimen from Key West and
designated it the lectotype (ANSP 35783), although the par-
tially disarticulated specimen from Cedar Keys (ANSP 35782)
probably is the one Pilsbry illustrated. Watters’ illustration of
very wide valves and his description of a striated jugum in-
dicate that the Key West lectotype and the Cedar Keys
specimen are conspecific.

The Cedar Keys specimen (Fig. 72), now a paralec-
totype, is broken into five pieces: valves i-iii, valves vi-vii, valve
vill, a broken intermediate valve (iv or v), and a fragment of
that valve imbedded in a piece of the girdle. Overall length
of the total specimen, estimated from its parts, is about 8 mm.
The strongly incised jugum demonstrates that the specimen
is conspecific with those reported as Acanthochitona pygmaea
herein.

Despite a great quantity of literature which states other-
wise, Acanthochitona pygmaea (Pilsbry, 1893) is not A.
spiculosa (Reeve, 1847). That conclusion is supported by
several observations: 1) there are no incisions on the jugum
of A. spiculosa; 2) intermediate valves of A. pygmaea are much
wider than long, whereas those of A. spiculosa are relatively
more narrow; 3) the syntypes of A. spicu/osa are considerably
larger than nearly all of the 924 intact A. pygmaea examined
herein; only two specimens of A. pygmaea were as large (21.0
mm, Turks and Caicos Ids., 21.2 mm, Puerto Rico) as the
smallest of the five syntypes (21.0-33.0 mm) of A. spiculosa.

Because this species is so common in Florida and the
northern Caribbean, most literature records of Acanthochitona
spiculosa actually represent A. pygmaea. Dall (1889a)
launched more than 90 years of taxonomic turmoil by in-
cluding Cedar Keys, west Florida, and the Florida Keys within
the range of A. spiculosa, indicating that his concept of A.
spiculosa included the species Pilsbry later described as A.
pygmaea. A. pygmaea is the only species of Acanthochitona
which occurs at Cedar Keys and nearshore west Florida.
Likewise, the A. spiculosa of Bermuda (Peile, 1926; Jensen
and Harasewych, 1986) is A. pygmaea. Among material | ex-
amined were specimens of A. pygmaea previously identified
as A. spiculosa by Kaas (RMNH), Watters (Bullock collection),
and Ferreira (CAS, IRCZM). Leloup (1941) recognized A.
pygmaea and illustrated valve viii of a specimen from Florida,
but specimens he reported from Venezuela and Colombia
could have been a new species described hereafter. Kaas
(1972) treated A. pygmaea and A. spiculosa separately, but
specimens he reported as A. spiculosa from Gulfport (RMNH
K3731) and Bonefish Key (RMNH K2852), Florida, are A.
pygmaea. It is doubtful that the specimens Kaas reported as

A. pygmaea were that species, as evidenced by his descrip-
tion of only 12-15 spicules in dorsal tufts and other features
more characteristic of several other species.

Where both species occur together in Florida and the
northern Caribbean, it is not uncommon to find specimens
of Acanthochitona andersoni in lots of A. pygmaea. Lots ex-
amined here that included both species are CAS 063321, col-
lected at Key West by Hemphill; ANSP 325864, a paratype
lot of A. andersoni Watters; and two unnumbered lots from
West Summerland Key in the Bullock collection.

Acanthochitona pygmaea is common in Florida, the
Bahama Islands, Yucatan, and Puerto Rico, but | have seen
no specimens southward from Saba Bank. In addition to
Leloup’s (1941) records from Venezuela and Colombia, A.
pygmaea has been reported from several locations in Brazil
by Righi (1971), who illustrated only the short dorsal spicules,
marginal spicules, and radula; those records need con-
firmation.

Acanthochitona venezuelana Lyons, sp. nov.
Figs. 73-80

TYPE MATERIAL: HOLOTYPE: Length approximately 20.0 mm
(curled), North of La Guardia, Isla de Margarita, Venezuela, 12 June
1987, C. Franz, collector, USNM 859317. PARATYPES: 4 spec., all
curled, approximately 16.0-19.0 mm, collected with holotype, FSBC
| 32569 (2), RMNH 55988 (1), Bullock collection (1).

TYPE LOCALITY: North of La Guardia, Isla de Margarita,
Venezuela.

DISTRIBUTION: Isla de Margarita, Venezuela.

DESCRIPTION: Largest specimen (holotype) approximately
20.0 mm long, 10.0 mm wide including girdle; valves occupy-
ing about 50% of total specimen width. Valves i-vii white with
scattered black maculations arranged in vaguely concentric
arcs anterior of beaks; jugum yellow-brown or mauve, usual-
ly with faint flush of mauve on tegmentum near beak. Valve
viii with black maculation covering most of tegmentum. Gir-
dle noticeably spiculose, tan to gray, with pale green spicules
in anterior and sutural dorsal tufts.

Valve i semilunate (Fig. 73), wider than long, posterior
margin straight, with anterior insertion plate bearing 5 U-
shaped slits; tegmentum occupying approximately 70% of
valve width. Valves ii-vii beaked posteriorly (Figs. 74, 75);
tegmentum oblate, about 1.6 times as wide as long, with con-
vex anterolateral margins; sutural laminae prominent, very
wide, broadly rounded anteriorly, separated by wide, U-shaped
sinus, with single deep slits along anterolateral margins. Valve
viii pentagonal (Fig. 76), widest anterolaterally, dropping away
rapidly behind posterior, elevated, prominently pointed mucro
(Fig. 77); sutural laminae well-developed, markedly concave
anteriorly, sharply produced at anterolateral corners; 2 nar-
row slits in posterior insertion plate.

Surface of jugum with smooth veneer overlying layer
of numerous thin, longitudinal striae; both layers fragile, easily
damaged, revealing honeycombed subjugal constructional
elements beneath. Tegmentum of all valves with flat, oval
pustules 220 um long, elongate near jugum, smaller (130 um),

LYONS: CARIBBEAN ACANTHOCHITONIDAE 97

Figs. 73-78. Acanthochitona venezuelana Lyons, sp. nov. Fig. 73. Valve i ex 19.0 mm paratype; Margarita Id., Venezuela; FSBC | 32569. Fig.
74. Valve iv, same specimen. Fig. 75. Valve v, same specimen. Fig. 76. Valve viii, same specimen. Fig. 77. Valve viii, 18.0 mm paratype;
same lot; lateral view. Fig. 78. Tegmental pustules, valve iv, same specimen (field width = 315 um).

more rounded, subspatulate near outer margins (Fig. 78);
macresthete subcentral, 5-8 micresthetes of nearly same
diameter as macresthete clustered mostly on adapical half
of pustule surface.

Girdle upper surface obviously spiculose, densely
covered with straight to slightly curved, sharp-tipped, clear,
glassy spicules (Figs. 79, 80), round in cross-section, about
300-600 um long, overlying and generally obscuring mat of
tiny (75 um) slender spicules. Dorsal spicules gradually in-
creasing in length to merge with marginal fringe, where they
are longest (about 1 mm); no demarcation or change in form
between dorsal and marginal spicules; 18 anterior and dor-
sal tufts with about 25 pale green, slender, straight, sharp-
pointed spicules up to 1.5 mm long; lower surface covered
with small (100 um), densely packed, straight, slender spicules

directed toward periphery.

DISCUSSION: Acanthochitona venezuelana most resembles
A. avicula (Carpenter, 1864). Watters (1981) noted the relation-
ship between the western Atlantic A. pygmaea (as A.
spiculosa) and the eastern Pacific A. avicula. Like A. pygmaea,
A. avicula has broad intermediate valves (Fig. 81), longitudinal
incisions on the jugum, and drop-shaped pustules. A.
venezuelana has broad valves with drop-shaped to spatulate
pustules but lacks jugal incisions. Most notably, dorsal girdle
spicules of A. avicula and A. venezuelana virtually are iden-
tical. The combination of high, pointed mucro, more narrow
anterior end of the jugum, and possession of mostly ovate
to subspatulate tegmental pustules separate A. venezuelana
from A. avicula.

98 AMER. MALAC. BULL. 6(1) (1988)

oS

Figs. 79, 80. Acanthochitona venezuelana Lyons, sp. nov. Fig. 79. Holotype, approximately 20.0 mm (curled), lateral view; Margarita Id., Venezuela;
USNM 859317. Fig. 80. Holotype, dorsal view. Fig. 81. Acanthochitona avicula (Carpenter, 1864); entire specimen, 12.4 mm; Puertocitos, Baja

California, Mexico; FSBC | 32570.

Acanthochitona avicula, A. pygmaea and A.
venezuelana join A. asterigera, A. hirudiniformis, and A. lineata
and A. hemphilli, A. rhodea, and A. ferreirai as groups with
one eastern Pacific and two western Atlantic species. Although
specimens of A. venezuelana have been seen only from
Margarita Island, the species probably has a wider distribu-
tion across the Caribbean coast of South America and could
replace A. pygmaea in that region. Dautzenberg (1900)
reported a curled specimen (2.5 x 2.5 mm) of A. pygmaea
dredged from 11 m at Los Testigos very near Isla Margarita,
and Leloup (1941) reported a curled specimen (3 x 2.5 mm) of
A. pygmaea dredged from 12-15 fm (22-27 m) off Cabo la Vela,
Colombia; a specimen from Florida, not the southern Carib-
bean, was illustrated by Leloup (his fig. 2, reproduced as figs.
82-84 by Kaas, 1972). Kaas (1972) reported no specimens of
A. pygmaea from farther south than St. Barts, Saba, and St.
Eustatius, and | have seen no A. pygmaea from any area south
of Saba Bank. Thus, it is possible that specimens reported
by Dautzenberg and by Leloup as A. pygmaea could have
been A. venezuelana.

ETYMOLOGY: Named for Venezuela, the Caribbean nation
where the specimens were collected.

Acanthochitona roseojugum Lyons, sp. nov.
Figs. 82-92

Acanthochitona pygmaea, Lyons, 1981: 36 (pars, Dry Tortugas
sta. 2 only) [non A. pygmaea (Pilsbry, 1893)].

TYPE MATERIAL: HOLOTYPE: Length 12.2 mm, width 6.0 mm,
Bartlett Hill, Eight Mile Rock, Grand Bahama Island, 0-0.5 m, 29 Aug
1984, W. G. Lyons, collector, USNM 859316. PARATYPES: FLORIDA:
4 spec., 8.1-8.7 mm, Bird Key Reef, Dry Tortugas, 0.5-1.0 m, 4 Oct
1979, FSBC | 32535. —1 spec., 6.4 mm, Florida Middle Ground,
28°35.0’N, 84914.9’W, 31 m, 19 May 1977, FSBC | 24598. —1 spec.,
10.6 mm, Peanut Id., Palm Beach Inlet, 0-1 m, 29 Aug 1982, FSBC
| 32536. BAHAMAS: 4 spec., 10.0-12.2 mm (2 curled), collected with
holotype, ANSP A12124 (1), RMNH 55989 (1), FSBC | 32537 (2). —1
spec., 12.4 mm, Caravel Beach, Freeport, Grand Bahama, 1 m, 30
Aug 1984, FSBC | 32538. :

OTHER MATERIAL EXAMINED: FLORIDA: 1 valve, Florida Mid-
dle Ground, 28°38.1’N, 84°16.3’W, bottom sediments, 28.6 m, 21 May
1977, FSBC | 32533. —8 valves, Florida Middle Ground, 28°35’N,
84°18’W, bottom sediments, 25.6-38.1 m, 7 Mar 1976, FSBC | 32532.
BAHAMAS: 4 valves, Gibson Cay, Andros, beach drift, 2 Sept 1971,
FSBC | 32531. HONDURAS: 1 spec., 16.2 mm, Utila Id., June 1987,
Sunderland collection.

TYPE LOCALITY: Bartlett Hill, Eight Mile Rock, Grand
Bahama Island.

DISTRIBUTION: Eastern Gulf of Mexico at Florida Middle
Ground to Dry Tortugas, southeast Florida, the Bahama
Islands, and Honduras; intertidal to 31 m.

DESCRIPTION: Largest specimen 16.2 mm long, 8.3 mm wide
including girdle; valves occupying 30-35% of total specimen
width (Figs. 82-84); tegmentum variously white with brown
flecks or pale pinkish white variegated with greenish black;

LYONS: CARIBBEAN ACANTHOCHITONIDAE

99

Figs. 82-84. Acanthochitona roseojugum Lyons, sp. nov. Fig. 82.
Holotype, 12.2 mm; Eight Mile Rock, Grand Bahama; USNM 859316.
Fig. 83. Paratype, 8.5 mm; Dry Tortugas, Florida; FSBC | 32535. Fig.
84. Paratype, 8.1 mm; same lot as 83.

jugum white or pink, suffused on some valves with bright rose
spots; girdle white or buff.

Valve i semilunate (Figs. 85, 86), wider than long,
margin straight posteriorly, with anterior insertion plate bear-
ing 5 U-shaped slits; tegmentum occupying 60-65% of valve
length. Valves ii-vii beaked posteriorly (Figs. 87, 88); tegmen-
tum subpentagonal, wider than long, with convex to slightly
sinuous anterolateral margins; sutural laminae large, flared
anterolaterally, with broadly rounded anterior tips separated
by broad, U-shaped sinus; single shallow slits along lateral
margins. Valve viii with tegmentum subovate (Figs. 89, 90),

Figs. 85-90. Acanthochitona roseojugum Lyons, sp. nov. Fig. 85. Valve
i ex 10.0 mm paratype; Eight Mile Rock, Grand Bahama; FSBC |
32537. Fig. 86. Valve i ex 8.2 mm paratype; Dry Tortugas, Florida;
FSBC | 32535. Fig. 87. Valve iv, same specimen as 85. Fig. 88. Valve
iv, same specimen as 86. Fig. 89. Valve viii, same specimen as 85.
Fig. 90. Valve viii, same specimen as 86.

widest between mucro and anterior margin; mucro elevated,
slightly posterior of center; sutural laminae large, broad, sub-
quadrate; 2 small slits in posterior insertion plate.

Jugum elevated, strongly demarked, smooth, narrow,
sides parallel, extending anteriorly beyond tegmental margin.
Tegmentum of all valves covered with subovate to spatulate,
flattened pustules (Figs. 91, 92) 120-140 um long, 80 um wide,
with single, subcentral macresthete, two pairs of micresthetes,
second pair near juncture of apex and tegmental plain.

Girdle upper surface covered with small (40 pm)
slender, sharp-tipped spicules; 18 anterior and sutural tufts
with 10-18 straight, relatively robust, vitreous spicules 1.25 mm
long, Surrounded by many similar but smaller (250 um)
spicules; marginal spicules sharp-tipped, vitreous, short (300
um) anteriorly and laterally, more than twice as long posterior-
ly; underside covered with fine (80 um), sharp-tipped, vitreous
spicules directed toward periphery.

DISCUSSION: Florida specimens generally have paler color
on the tegmentum and girdle, and valves seem to be slightly
more protracted. However, the rose spots, extended jugum,
and tegmental pustule morphology indicate that Bahamian
and Florida populations are conspecific.

Intact specimens of Acanthochitona roseojugum hardly
seem separable from A. andersoni Watters, 1981. Differences
useful to sort specimens are almost subjective. Intermediate
valves of A. roseojugum are wider and more flattened anterior-
ly, whereas those of A. andersoni are more narrow and arched.
The jugum of A. roseojugum is separated more distinctly from
the tegmentum than is that of A. andersoni. Rose-colored
spots occur on all or part of the jugum of at least valve iii

100 AMER. MALAC. BULL. 6(1) (1988)

Figs. 91, 92. Acanthochitona roseojugum Lyons, sp. nov., tegmental
pustules (field widths = 385 um). Fig. 91. Bahamas; same specimen
as 85. Fig. 92. Florida; same specimen as 86.

of A. roseojugum and sometimes occur on the jugum of all
intermediate valves (ii-vii); sutural laminae and undersides of
all valves are pink. | have seen two entirely rose-colored
specimens of A. andersoni, but those specimens were
distinguishable by their highly arched, more narrow in-
termediate valves. Some specimens of A. pygmaea from the
Bahamas and Puerto Rico are flushed with pale pink on some
intermediate valves, but these are immediately separated from
A. roseojugum by strongly incised grooves on the jugum, wider
tegmentum on intermediate valves, smaller sutural laminae,
and many green spicules rather than few white spicules in
the anterior and sutural tufts.

Any resemblance of Acanthochitona roseojugum to A.
andersoni and A. pygmaea is disspelled by inspection of disar-
ticulated valves. The proportionately large insertion plate and
small tegmentum of valve i, flared sutural laminae and ex-

tended, strongly demarked, smooth jugum of valves ii-viii, and
small slits of valve viii all resemble features of species in the
A. hemphilli complex. However, the straight posterior margin
of valve i and the girdle species of A. roseojugum differ con-
siderably from those of species in the A. hemphilli complex.

The additional asymmetrical slits on insertion plates
of valves i and viii of the illustrated Bahamian specimen (Figs.
85, 89) represent anomalies that occur occasionally in many
species of Acanthochitona.

ETYMOLOGY: From Latin ‘‘roseus’’, rose-colored, and
“jugum’’, a ridge (i.e. jugum).

Acanthochitona balesae Abbott, 1954
Figs. 93-104

Acanthochitona balesae Pilsbry, 1940: pl. 12, fig. 5 (nomen
nudum). Abbott, 1954: 318; 1974: 406. Watters, 1981:
175, 176, pl. 3, figs. a-c.

Acanthochitona elongata Kaas, 1972: 51-53, figs. 90-94,
pl. 2, fig. 3. Ferreira, 1985: 212.

Acanthochitona interfissa Kaas, 1972: 53-55, figs. 95-107.

Choneplax lata, Ferreira, 1985: 208-213 (pars) [non Choneplax
lata (Guilding, 1829)].

TYPE MATERIAL: HOLOTYPE: A. balesae: ANSP 349331 (not ex-
amined). A. interfissa: 5.5 mm; Monos, Avalon Bay, Trinidad; 10 Jan
1955; RMNH 9092. PARATYPES: A. interfissa: TRINIDAD: 1 spec.,
7.0 mm; collected with holotype; RMNH 9093. ARUBA: 5 disar-
ticulated intermediate valves; Malmok, Arasji; 14 Aug 1955; RMNH
4502. —1 spec., 8.8 mm; same locality and date; RMNH 9094.

OTHER MATERIAL EXAMINED: FLORIDA: 2 spec., 6.7, 9.6 mm,
north side Vaca Key, 0-1 m, 1 Oct 1979, FSBC | 32558. —1 spec.,
curled, same locality, 4 Aug 1980, FSBC | 32571. —1 spec., 9.2 mm,
Bonefish Key, CAS 063327. —1 spec., 9.3mm, Peanut Id., Palm Beach
Inlet, 0-1 m, 17 Aug 1982, FSBC | 30761. —2 spec., 4.4, 68 mm, 3
km south of St. Lucie Inlet, 2-3 m, 18 May 1978, IRCZM 61:008. —1
spec., 3.7 mm, same location and date, IRCZM 61:007. BAHAMAS:
1 spec., 8.4 mm, Eight Mile Rock, Grand Bahama, 0.5-1.0 m, 21-23
May 1981, FSBC | 32559. —2 spec., 9.5, 10.6 mm, Bartlett Hill, Eight
Mile Rock, 0-0.5 m, 29 Aug 1984, FSBC | 32040. JAMAICA: 1 in-
termediate valve, Drunkeman’s Key, RMNH. ST. EUSTATIUS: 8 spec.,
2.4-4.4 mm, Tumble Down Dick Bay, RMNH. TRINIDAD: See type
material. VENEZUELA: 1 spec., 7.8 mm, Tortuga Id., CAS 063326.
ARUBA: 3 spec., 2.5-8.3 mm, Malmok, 14 Aug 1955, RMNH. —1
spec., 7.0 mm, Seroe Colorado, 2 May 1955, RMNH. —1 spec., 5.1
mm, Rincon, 7 May 1955, RMNH. See also type material. PANAMA:
9 spec., 2.0-4.0 mm, Galeta Id., Canal Zone, Bullock collection. —10
spec., 3.0-5.0 mm, Galeta Id., Bullock collection. —10 spec., 3.0-7.0
mm, Galeta Id., Bullock collection.

TYPE LOCALITY: Bonefish Key (= Fat Deer Key, between
Vaca Key and Crawl Key, Monroe County, Florida; see Kaas,
1972) (original designation).

DISTRIBUTION: South Florida and Grand Bahama Island to
Caribbean coast of Panama and Trinidad.

DESCRIPTION: Largest specimen 10.6 mm long, 3.7 mm wide
including girdle; valves occupying about 33% total specimen
width (Figs. 93, 94). Exposed valves white, usually with beige,
olive, or brown maculations, occasionally some valves entirely
brown-black; intermediate valves noticeably longer than wide.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 101

Figs. 93-99. Acanthochitona balesae Abbott, 1954. Fig. 93. Whole specimen, 9.6 mm; Vaca Key, Monroe County, Florida; FSBC | 32558. Fig.
94. Entire specimen, 6.7 mm; same lot. Fig. 95. Valve i ex curled specimen; Vaca Key, Florida; FSBC | 32571. Fig. 96. Valve iv, same specimen.
Fig. 97. Valve v, same specimen. Fig. 98. Valve vili, same specimen. Fig. 99. Tegmental pustules, valve iv, same specimen (field width = 240 ym).

Girdle beige to tan (bleached totally white in some preserved
specimens), with green, brown, or black patches between
white spicule clusters of dorsal tufts.

Valve i semilunate (Fig. 95), slightly wider than long,
markedly concave posteriorly, with anterior insertion plate
bearing 5 slits; tegmentum occupying about 90% total valve
length. Posterior margins of valves iii-vi strongly produced
(Figs. 96, 97), those of remaining valves nearly straight;
tegmentum longer than wide, subpentagonal, widest at
posterolateral corners, with straight anterolateral margins;
sutural laminae long, narrow, separated at anterior, acute tips
by U-shaped sinus, margins parallel with longitudinal axis of
valves, with or without single, narrow slits along margins. Valve

viii about as wide as long (Fig. 98), rounded posteriorly, with
mucro posterior of center; tegmentum subpentagonal, longer
than wide, dropping rapidly behind mucro; sutural laminae
long, narrow, with straight anterolateral margins, subacute
anterior tips separated by U-shaped sinus; 2 small slits in
posterior insertion plate.

Jugum moderately expanded anteriorly, smooth, with
irregular lateral margins merging with tegmental pustules.
Tegmentum of all valves with peg-like, elevated, ovate to
spatulate pustules (Fig. 99) about 90 um long, 45 nm wide,
with single subcentral macresthete, usually 3-4 micresthetes.

Girdle upper surface evenly covered with short (80 um),
straight to slightly bent, blunt or sharp-tipped, light or dark

102 AMER. MALAC. BULL. 6(1) (1988)

Figs. 100-104. Acanthochitona balesae Abbott, 1954. Intermediate
valves of disarticulated paratype of A. interfissa Kaas, 1972; Malmok,
Arasji, Aruba; RMNH 4502. Length of largest valve (Fig. 100) 1.6 mm,
including sutural laminae.

colored spicules; 18 anterior and sutural tufts comprised of
about 50 straight, slender, sharp-tipped, vitreous spicules up
to 700 nm long; margin fringed with straight, slender, sharp-
tipped spicules 250-280 nm long; underside evenly covered
with short (50-60 ym), straight, sharp-tipped spicules directed
toward periphery.

DISCUSSION: Several names have been proposed for this
species. Pilsbry (1940) illustrated without text a chiton he
called Acanthochitona balesae from Bonefish Key, Florida,
thereby creating a nomen nudum. Abbott (1954) included A.
balesae ‘Pilsbry 1940’ from Bonefish Key, with brief differen-
tial diagnostic remarks. Kaas (1972) recognized the nude
status of Pilsbry’s name; to rectify that problem, he described
four specimens from Bonefish Key (RMNH) and named them
A. elongata. In the same paper, Kaas named A. interfissa from
Trinidad and Aruba and noted similarities between that
species and A. elongata. Abbott (1974) included A. balesae
‘Pilsbry’ Abbott, repeated his diagnostic comments, and stated
that A. elongata was a synonym. Bullock (1974) pointed out
that Kaas ‘‘overlooked the fact that Abbott ... validated
Pilsbry’s name, and A. elongata Kaas must be considered a
junior synonym of A. balesae ‘Pilsbry’ Abbott.’ Bullock also
remarked that the relationship between A. interfissa and A.
balesae should be investigated. Watters (1981) relegated both
A. elongata and A. interfissa to the synonymy of A. balesae
and designated a lectotype (ANSP 349331; Bonefish Key) for
A. balesae. Ferreira (1985) incorrectly stated that Abbott (1974)
regarded A. balesae to be asynonym of A. elongata. Ferreira
clearly considered Abbott’s diagnosis inadequate and without
priority over A. elongata. He agreed with Watters that A. in-
terfissa is a synonym of A. elongata, but he also relegated

A. andersoni Watters, 1981, to the synonomy of A. elongata.
Finally, Ferreira declared all the above taxa to be juveniles
and secondary synonyms of Choneplax lata (Guilding, 1829).

The International Code of Zoological Nomenclature re-
quires that, to be available, a species name introduced after
1930 must be accompanied by a description or definition that
states in words characters that are purported to differentiate
the taxon [Article 13(a)(i); ICZN, 1985]. Abbott’s (1954) account
of Acanthochitona balesae, although brief, addressed size,
proportions, pustule morphology, shape and ornamentation
of the jugum, and a location where the species occurs; some
characters were compared with those of A. pygmaea. Such
treatment satisfies the requirements of ICZN Article 13, so A.
balesae Abbott, 1954, is valid, and A. elongata Kaas, 1972,
is a junior synonym.

| examined the holotype and three of the four paratypes
of Acanthochitona interfissa Kaas and the holotype and seven
paratypes of A. andersoni Watters. | found no characters upon
which to separate the holotype and paratypes of A. interfissa
from topotypic specimens of A. balesae from Bonefish Key,
so | cannot refute contentions by Watters (1981) and Ferreira
(1985) that A. interfissa is a synonym of A. balesae. However,
A. andersoni is not a synonym of A. balesae, and neither name
is a synonym of Choneplax lata.

Several problems are associated with the original
description and type series of Acanthochitona interfissa. Kaas
reported the holotype and a paratype from Trinidad and three
paratypes from Aruba. He reported that he disarticulated and
illustrated the paratype from Trinidad. However, although the
valves and spicules of that specimen now are almost totally
dissolved in preservative, the specimen is intact, as is the
holotype. One of the Aruba paratypes has been disarticulated.
Five of the valves remain.(Figs. 100-104), but valves i, viii, and
an intermediate valve are missing; none of the valves
resembles the curiously misshapen valve ii illustrated by
Kaas.

Except for valve viii, the description and illustrations
of valves of Acanthochitona interfissa (Kaas, 1972: figs. 95-101)
seem indistinguishable from those of A. balesae. Valve viii
of A. interfissa as illustrated by Kaas (his figs. 95-97) differs
from the corresponding valve of A. elongata (= A. balesae)
(Kaas, 1972: figs. 90, 91) by tegmental shape, pustule con-
figuration and size, jugal length and expansion, by posses-
sion of a greatly flared insertion plate and laminae, and most
notably, by possession of a medial third slit in the posterior
insertion plate. Conversely, valves i, ii, and iv of A. interfissa
(Kaas, 1972: figs. 98-101) are indistinguishable from those of
A. balesae whose corresponding valves Kaas described but
did not illustrate in the account of A. elongata which im-
mediately preceded that of A. interfissa.

Ferreira (1985) could have been prompted to combine
Acanthochitona andersoni with A. interfissa because of Kaas’
description of valve viii of the latter. Among the Caribbean
Acanthochitona species, valve viii of A. interfissa as illustrated
by Kaas most resembles that of A. andersoni, if the third slit
of A. interfissa is ignored. | found a single specimen of A.
andersoni among three A. ba/esae in an uncatalogued lot
(RMNH) from Malmok, Aruba, collected on the same date as

LYONS: CARIBBEAN ACANTHOCHITONIDAE 103

were the paratypes of A. interfissa. However, no species of
Acanthochitona normally possesses a third slit in valve viii.
Because the 3-slitted valve no longer accompanies the type
material, it seems best to regard the third slit as an anomalous,
additional one of the kind that sometimes occurs on other nor-
mally 2-slitted species.

Kaas (1972) described the sutural laminae of in-
termediate valves of Acanthochitona elongata as ‘‘unslit, but
with little excavations where the slits might be expected’’; for
A. interfissa, he described ‘‘valves with 1 slit, except valves
iv-vi which are unslit’” The specimen of A. balesae | dissected,
collected within 1 km of the type-locality, has distinct slits on
valves ii and vii but lacks slits on valves iii-vi. Kaas also
described a longitudinally striate jugum for A. elongata, which
he contrasted with the smooth jugum of A. interfissa. Although
longitudinal striae were sometimes visible beneath the sur-
face, | saw only a smooth jugum on all specimens of A.
balesae | examined.

Despite my inability to find objective differences be-
tween the two taxa, it should be noted that specimens of the
northern Caribbean Acanthochitona balesae and those of the
southern A. interfissa can be sorted by seemingly subjective
characters. Basically, southern specimens are smaller, more
drab, and have finer spicules and sculpture than northern
specimens. Using those ‘‘characters’’, all Florida and Baha-
mian specimens are assignable to A. balesae and all
specimens from St. Eustatius to Trinidad, Venezuela, Aruba
and Panama are assignable to A. interfissa. Further work may
yet reveal objective characters which can be used to
demonstrate two species within the group.

Watters’ (1981) drawings of valves from Puerto Rico are
too schematic to reveal with certainty whether they belong
to A. balesae.

Acanthochitona andersoni Watters, 1981
Figs. 105-109

Acanthochitona andersoni Watters, 1981: 173-176, pl. 2e-g,
pl. 4i.

Acanthochitona pygmaea, Lyons, 1981: 36 (pars, Dry Tortugas
Sta. 4 only) [non A. pygmaea (Pilsbry, 1893)].
Choneplax lata, Ferreira, 1985: 208-213 (pars) [non C. lata

(Guilding, 1829)].

TYPE MATERIAL: HOLOTYPE: 11.3 mm, Calliagua, St. Vincent,
Feb 1972, ANSP 332171. PARATYPES: FLORIDA: 1 spec., 6.4 mm,
off Destin, 55 m, ANSP 220834. —2 spec., 5.7, 7.5 mm, West Sum-
merland Key, Oct 1973, Bullock collection. —1 spec., curled, West
Summerland Key, 1 June 1974, Bullock collection. —1 spec., 5.3 mm,
off Boynton, 55 m, ANSP 220833. BAHAMAS: 1 spec., 9.5 mm, west
of Haulover, North Bimini, ANSP 325808. —1 spec., 7.5 mm, east of
Turtle Rocks, 6 m, ANSP 325864.

OTHER MATERIAL EXAMINED: FLORIDA: 1 spec., 5.7 mm,
Garden Key, Dry Tortugas, 0-2 m, 5 Oct 1979, FSBC | 32551. —3 spec.,
46-75 mm, Garden Key, 30 Apr 1975, CAS 063329. —1 spec., 63
mm, Key West, CAS 063321. —1 spec., 11.5 mm, West Summerland
Key, 1976, Bullock collection. —1 spec., curled, West Summerland
Key, 1978, Bullock collection. —1 spec., 8.0 mm, Missouri Key, 0.5-1.0
m, 25 July 1987, FSBC | 32557. —1 spec., curled, Burnt Point, Crawl
Key, 2.5 m, 4 Aug 1982, FSBC | 32426. —1 spec., 4.7 mm, Tennessee

Reef, off Long Key, 13.7 m, 12 July 1986, FSBC | 32556. —1 spec.,
7.0 mm, Elbow Reef, 25°07.7’N, 80°15.9’W, 18.3 m, 7 June 1979, IRCZM
61:018. —3 spec., curled, east of Elliott Key, RMNH. —1 spec., 8.8
mm, Peanut Id., Palm Beach Inlet, 0-1 m, 29 Aug 1982, FSBC | 30762.
BAHAMAS: 2 spec., 3.4, 10.1 mm, Bartlett Hill, Eight Mile Rock, Grand
Bahama, 0-0.5 m, 29 Aug 1984, FSBC | 32553. —1 spec., 7.5 mm,
Tamarind Beach Reef, Grand Bahama, 18 m, 28 Aug 1984, FSBC
| 32552. —2 spec., 7.2, 8.0 mm, Green Turtle Cay, Abaco, 0.5 m, May
1978, FSBC | 32550. PUERTO RICO: 1 spec., 6.0 mm, Isla Turramote,
La Parguera, May 1985, FSBC | 32554. SABA: 5 spec., curled, Fort
Bay pier, 7 July 1973, RMNH. ST. LUCIA: 2 spec., 8.0, 9.0 mm, Anse
Chastenet, 1-3 m, 4 Nov 1984, Bullock collection (1), FSBC | 32572
(1). ARUBA: 1 spec., 3.0 mm, Malmok, Arasji, 14 Aug 1955, RMNH.
BONAIRE: 1 spec., 9.0 mm, 2 km north of Kralendijk, 4m, 7 Oct 1986,
FSBC | 32555. CURAQGAO: 1 spec. (7), 2.0 mm, Piscadera Baai,
0-4 m, Apr 1966, Bullock collection. —1 spec. (?), 2.5 mm, Knip Baai,
6 Feb 1949, RMNH. VENEZUELA: 1 spec., crushed, Tortuga Id., 1
Aug 1936, RMNH.

TYPE LOCALITY: Calliagua, St. Vincent (original designation).

DISTRIBUTION: Both coasts of Florida, the Bahama Islands,
the Lesser Antilles, southern Netherlands Antilles, and
Venezuela. Watters (1981) also reported specimens from
Quintana Roo, Mexico, and Caribbean Panama.

DESCRIPTION: Largest specimen (holotype) 11.3 mm long,
48 mm wide including girdle; valves occupying about 50%
of total specimen width (Fig. 105). Exposed parts of valves
of holotype white, extensively mottled with black; most other
specimens white or light green with few brown or black flecks,
few specimens apricot or rose. Girdle white, buff, tan, or dark
brown, often with bar-like maculations; spicules translucent
white.

Valve i semilunate (Fig. 106), wider than long, slightly
to markedly concave posteriorly, with anterior insertion plate
bearing 5 slits; tegmentum occupying 70-75% total valve
length. Valves ii-vii prominently beaked posteriorly (Fig. 107);
tegmentum pentagonal, as wide or wider than long, with
slightly convex anterolateral margins; sutural laminae
moderately to considerably produced anteriorly, with vague
to distinct anterolateral angle, subacutely rounded anterior-
ly, separated by wide anterior sinus; single slits along lateral
margins. Valve viii pentagonal (Fig. 108), widest at
anterolateral corners, dropping away rapidly behind elevated,
postcentral mucro; sutural laminae well-developed, with
straight margins and sharply angled corners; 2 narrow,
relatively small slits in posterior insertion plate.

Jugum smooth, narrow, little expanded anteriorly,
merging laterally with tegmental pustules. Tegmentum of all
valves covered with ovate or subspatulate pustules (Fig. 109)
90-130 »m long, 60-80 um wide, with single adapical
macresthete, 2-6 micresthetes between macresthete and
apex.

Girdle upper surface covered with dense mat of very
small (40 um) slender spicules; 18 anterior and sutural tufts
comprised of 12-20 stout, straight, sharp-tipped vitreous
spicules up to 1.2 mm long, accompanied at base by many
sharp, slender, needle-like spicules about 200 um long; margin
fringed with stout, straight to slightly curved vitreous spicules
about 140 um long, with markedly larger (200 »m) but other-

104 AMER. MALAC. BULL. 6(1) (1988)

Figs. 105-109. Acanthochitona andersoni Watters, 1981. Fig. 105. Holotype, 11.3 mm; Calliagua, St. Vincent; ANSP 332171. Fig. 106. Valve
i ex 8.0 mm specimen; Anse Chastenet, St. Lucia; FSBC | 32572. Fig. 107. Valve iv, same specimen. Fig. 108. Valve viii, same specimen.
Fig. 109. Tegmental pustules, valve iv ex 8.0 mm specimen; Green Turtle Cay, Abaco, Bahamas; FSBC | 32550 (field width = 365 um).

wise similar spicules sparsely scattered throughout; under-
side covered with slender, sharp-tipped, vitreous spicules
about 80 um long directed toward periphery.

DISCUSSION: Specimens of Acanthochitona andersoni have
been confused with A. pygmaea, A. balesae, and Choneplax
lata. The smooth, not incised jugum and relatively narrow, not
widely rectangular intermediate valves distinguish A. ander-
soni from A. pygmaea. The tegmentum of intermediate valves
of A. andersoni is as wide or slightly wider than long, whereas
that of A. balesae is longer than wide. Morphology of tegmen-
tal pustules is also distinctive for each of the three species.
A. andersoni is not C. lata, as evidenced by possession of 2
distinct slits on valve viii. Ferreira (1985) identified lots CAS
063329 from Dry Tortugas and IRCZM 61:108 from Elbow Reef
as Choneplax lata and CAS 063321 from Key West as Acantho-

chitona spiculosa.

Acanthochitona bonairensis Kaas, 1972
Figs. 110-113

Acanthochitona bonairensis Kaas, 1972: 44, 45, figs. 72, 73,
pl. 3, figs. 1, 2. Ferreira, 1985: 207, 214.

Acanthochitona communis, Watters, 1981: 173.

Acanthochitona fascicularis, Kaas, 1985: 586.

TYPE MATERIAL: HOLOTYPE: 33 mm x 22 mm, Bonaire, RMNH.

DISCUSSION: Nothing can be added to the original descrip-
tion. Kaas (1972) noted the similarity in valve morphology be-
tween Acanthochitona bonairensis and the European species
A. communis (Risso, 1826), but also described considerably
shorter, more delicate girdle spicules on A. bonairensis than

LYONS: CARIBBEAN ACANTHOCHITONIDAE 105

~
x

“112

Figs. 110-112. Acanthochitona bonairensis Kaas, 1972. Fig. 110.
Holotype, 33.0 mm; Punt Vierkant, Bonaire; RMNH. Fig. 111. Valve
vii of holotype. Fig. 112. Valve viii of holotype.

Fig. 113. Acanthochitona bonairensis Kaas, 1972. Valve viii of holotype.
Fig. 114. Acanthochitona fascicularis (Linné, 1767). Valve viii ex
specimen from Roscoff, France; FSBC | 32427. Compare outline of
tegmentum with that of specimen in Fig. 113.

on A. communis. Watters (1981) ignored the described dif-
ferences and declared A. bonairensis to be a synonym of A.
communis. Kaas (1985) followed that synonymy in his review
of A. fascicularis (Linné, 1767), a senior synonym of A. com-
munis. However, Ferreira (1985) retained A. bonairensis as one
of the few Caribbean species he considered distinct.

| compared the holotype of Acanthochitona bonairen-
sis (Figs. 110-112) with specimens of A. fascicularis from
Roscoff, France (FSBC | 32427). Differences in valve mor-
phology (Figs. 113, 114) noted by Kaas (1972), although sub-
tle, were confirmed, as were marked differences in girdle
spicules. A. bonairensis remains known only from the
holotype. Discovery of more Caribbean specimens would help
considerably in interpretation of differences noted to date. Until
such specimens are found, | believe the differences in girdle
spicules provide sufficient reason to maintain A. bonairensis
as a Caribbean species distinct from the European A.
fascicularis.

Acanthochitona zebra Lyons, sp. nov.
Figs. 115-127

(?) Choneplax lata, Kaas, 1972: 55-58, figs. 108-116, pl. 2,
fig. 4 (pars) [non C. lata (Guilding, 1829)].

Acanthochitona sp. Lyons, 1981: 35, 36.

Choneplax lata, Ferreira, 1985: 208-213 (pars). [non C. lata
(Guilding, 1829)].

TYPE MATERIAL: HOLOTYPE: Length 15.0 mm, Silver Cove Canal,
Freeport, Grand Bahama Island, 0.5-1.5 m, 28 Aug 1984, W. G. Lyons,
collector, USNM 859319. PARATYPES: FLORIDA: 1 spec., 12.0 mm,
Long Key Reef, Dry Tortugas, intertidal, 11-12 May 1979, FSBC | 32479.
—6 spec., 7.0-11.2 mm, patch reef near Long Key Reef, Dry Tortugas,
1.5-2.5 m, 11-12 May 1979, ANSP A12125 (1), FSBC | 32478 (5). —1
spec., 4.5 mm, Tennessee Reef, off Long Key, 13.7 m, 12 July 1986,
FSBC | 32485. BAHAMAS: 1 spec., 11.3 mm, same locality and date
as holotype, FSBC | 32483. —1 spec., 9.7 mm, Caravel Beach,
Freeport, Grand Bahama, 1 m, Jan 1981, FSBC | 32480. —6 spec.,
5.0-10.0 mm, Tamarind Beach Reef, Grand Bahama, 18 m, 28 Aug
1984, RMNH 55990 (1), FSBC | 32482 (5). — 1 spec., 8.2 mm, Salt
Pond, Long Island, Aug 1975, CAS 063328. PUERTO RICO: 2 spec.,
7.2, 9.3 mm, Isla Turramote, La Parguera, 9.1 m, May 1985, FSBC
| 32484. BELIZE: 1 spec., 15.0 mm, Carrie Bow Cay, 0-1 m, 23 Mar
1981, IRCZM 61:092.

OTHER MATERIAL EXAMINED: FLORIDA: 1 spec., 3.4 mm, east
of Elliott Key, 2-6 m, 5 Sept 1963, RMNH. BAHAMAS: 7 intermediate
valves, Gold Rock, Grand Bahama, bottom sediments, 24.4 m, May-
July 1981, FSBC | 32481. —7 intermediate valves, Grand Bahama,
bottom sediments, May 1981, R. Quigley collection. ARUBA: 2 spec.
(?), both small, missing valve viii, Paardenbaai rif, 28 Apr 1955, RMNH.
CURAGAO: 3 spec., 5.2-7.3 mm, Piscadera Baai, 27 July 1973, RMNH.
—2 spec. (?), 2.7, 2.9 mm, Caracas Baai, 22 Apr 1955, RMNH. —1
spec., 6.5 mm, Spaanse Water, 17 Nov 1968, RMNH. —1 spec. (7),
3.4 mm, Awa di Oostpunt, 0.25-1.0 m, 22 Feb 1970, RMNH.

TYPE LOCALITY: Silver Cove Canal, Freeport, Grand
Bahama Island.

DISTRIBUTION: Dry Tortugas, Florida Keys, and Grand
Bahama Island to Puerto Rico and Belize, Aruba and Curagao;
intertidal to 18 m, single valves from sediments in 24.4 m.

106 AMER. MALAC. BULL. 6(1) (1988)

Figs. 115-117. Acanthochitona zebra Lyons, sp. nov. Fig. 115. Holotype,
15.0 mm; Freeport, Grand Bahama; USNM 859319. Fig. 116. Paratype,
8.3 mm; Tamarind Beach Reef, Grand Bahama; FSBC | 32482. Fig.
117. Paratype, 11.1 mm; Dry Tortugas, Florida; FSBC | 32478.

DESCRIPTION: Largest specimen (holotype) 15.0 mm long,
7.2 mm wide including girdle; valves and girdle occupying ap-
proximately equal portions of total specimen width (Figs.
115-117). Valve i with 3-5 olivaceous or brown concentric
bands, expressed on valves ii-vii as transverse stripes
(chevrons) extending posterolaterally from jugum; bands
usually strongest on valves i-v, commonly obscured by overall
dark olive or brown color on valves iv and vii; valve viii most-
ly white, with single large olivaceous spots on lateral areas.
Girdle white with irregular olivaceous or green bands cross-

ing upper surface from valves to peripheral margins,
sometimes with broad, black spots at middle or elsewhere
on each side.

Valve i semilunate (Fig. 118), wider than long, posterior
margin straight, slightly beaked, with anterior insertion plate
bearing 5 slits; tegmentum occupying 80-85% of valve length.
Valves ii-vii strongly beaked posteriorly (Fig. 119); tegmentum
evenly to broadly pentagonal, with convex anterolateral
margins; sutural laminae moderately narrow, curving
anteromedially from posterolateral corners of tegmentum, with
subacute anterior tips separated by broad sinus of same width
as anterior end of jugum; single narrow slits along lateral
margins. Valve viii tegmentum roughly ovate, widest mesial-
ly, truncate anteriorly, extending to overhang posterior edge
of insertion plate (Fig. 120). Mucro distinctly posterior; sutural
laminae extending obliquely anteriorly, subquadrate, of
moderate length; two slits in posterior insertion plate very fine,
barely discernible with dissecting microscope. Valve mor-
phology of Puerto Rican juveniles and Floridan adults as il-
lustrated (Figs. 121-126).

Jugum of valves ii-viii smooth, wedge-shaped, widest
anteriorly. Tegmentum covered with densely packed, flattened,
spatulate pustules (Fig. 127), approximately 80-100 nm long,
70 wm wide, radiating anteriorly from beak of valve i,
anterolaterally from jugum of valves ii-vii, and from mucro of
valve viii; pustules with single macresthete near apex, 4-7
micresthetes surrounding macresthetes, sometimes more on
Florida specimens; many additional micresthetes dispersed
across surface of tegmental plain.

Girdle upper surface covered with fine (100 pm)
spicules; 18 anterior and sutural tufts comprised of 8-10 red-
dish brown, amber, or white, moderately long (to 650 um),
slightly curved, blunt-tipped spicules; marginal spicules
straight or slightly curved, approximately 550 um long, with
blunt tips, white, sometimes alternating with amber; under-
side covered with fine (60 um), sharp-tipped spicules directed
toward periphery.

DISCUSSION: The olivaceous stripes on the tegmentum of
Acanthochitona zebra strongly resemble those of A. /ineata,
and A. astrigera sometimes has white stripes or maculations
on the dark blue-green tegmentum of some valves. Moreover,
all three species occurred together at the type-locality of A.
zebra. However, A. zebra can be separated readily from the
other two species by its extremely posterior mucro, from which
the tegmentum drops rapidly to overhang the posterior inser-
tion plate, and by the dorsal tufts of the girdle, which contain
only 8-10 blunt-tipped spicules. Pustular shape, as well as
location of macrestheses and micresthetes, further distinguish
A. zebra from A. astrigera and A. lineata.

Valve proportions of Florida specimens differ somewhat
from those of specimens from the Bahamas and Puerto Rico,
but morphology of valve viii and the tegmental pustules, as
well as the color pattern, indicate they are conspecific. Five
RMNH lots from Aruba and Curagao appear to be this species,
but the concentric bands and stripes are only weakly ex-
pressed on the four largest (5.2-7.3 mm) specimens and are
not evident at all on the five smaller (2.7-3.4 mm) specimens.

LYONS: CARIBBEAN ACANTHOCHITONIDAE 107

Figs. 118-127. Acanthochitona zebra Lyons, sp. nov. Fig. 118. Valve i ex 10.0 mm paratype; Tamarind Beach Reef, Grand Bahama; FSBC |
32482. Fig. 119. Valve iv, same specimen. Fig. 120. Valve viii, same specimen; ventral view showing underhung posterior insertion plate with
vestigial slits. Fig. 121. Valve i ex 7.2 mm paratype; Isla Turramote, Puerto Rico; FSBC | 32484. Fig. 122. Valve iv, same specimen. Fig. 123.
Valve viii, same specimen. Fig. 124. Valve i ex 11.0 mm paratype; Dry Tortugas, Florida; FSBC | 32478. Fig. 125. Valve iv, same
specimen. Fig. 126. Valve viii, same specimen. Fig. 127. Tegmental pustules, valve iv, same specimen as 118 (field width = 335 pm).

Ferreira (1985) identified the CAS specimen from Long
Island, Bahamas, and the IRCZM specimen from Carrie Bow
Cay, Belize, as Choneplax lata.

ETYMOLOGY: From the Amharic ‘‘zebra’’, as in Equus zebra,
an African equine with similar markings.

Genus Choneplax Dall, 1882
Choneplax lata (Guilding, 1829)
Figs. 128-145

Chitonellus latus Guilding, 1829: 28.

Chiton strigatus Sowerby, 1840: 289.

(?)Chiton hastatus Sowerby, 1840: 290, pl. 16, fig. 4.

Choneplax latus, Pilsbry, 1893: 60, pl. 8, fig. 15.

Choneplax lata, Kaas, 1972: 55-58, figs. 108-116, pl. 2, fig. 4
(pars). Ferreira, 1985: 208-213 (pars).

MATERIAL: BAHAMAS: 4 spec., large, curled, West End, Grand
Bahama, intertidal, May 1977, FSBC | 32546. —3 spec., 17.7-22.4 mm,
Settlement Point, West End, Grand Bahama, 2 m, 23 May 1981, FSBC

108 AMER. MALAC. BULL. 6(1) (1988)

| 32547. —55 spec., 6.5-32.0 mm, Bahama Beach Canal, West End,
Grand Bahama, intertidal, 29 Aug 1984, FSBC | 32548. —1 spec.,
26.0 mm, New Providence, CAS 063325. —1 spec., 15.0 mm, Nicolls
Town, Andros, 2 m, July 1976, CAS 063323. CUBA: 2 spec., 15.9, 17.0
mm, Phillips Park, Guantanamo Bay, intertidal, 9 Apr 1984, FSBC
| 32549. BELIZE: 3 spec., 19.0-22.0 mm, Carrie Bow Cay, 0-1 m, 23
Mar 1981, IRCZM 61:051. —1 spec., 9.0 mm, same locality and date,
IRCZM 61:053. HONDURAS: 1 spec., 10.0 mm, First Bight, Roatan,
1-2 m, Aug 1982, FSBC | 32073. GUADELOUPE: 4 spec., 13.0-20.0
mm, Guadeloupe, 28 May 1978, CAS 063324.

TYPE LOCALITY: St. Vincent (original designation).

DISTRIBUTION: Grand Bahama Island, Cuba, Belize, Hon-
duras, Guadeloupe, St. Vincent; intertidal and shallow (1-2 m)

subtidal zones. Kaas (1972) reported specimens from the
Virgin Islands, Tobago, Bonaire, and Curagao.

DESCRIPTION: Largest specimen 32.0 mm long, 13.7 mm
wide including girdle; valves occupying approximately 33%
of total specimen width (Fig. 128), proportionally more in
juveniles (Fig. 129). Valves brown-black, frequently eroded
to create bluish white bands between jugum and lateral
margins. Girdle yellow to greenish gold, often with brown or
black band across middle.

Valve i semilunate (Fig. 130), wider than long, slightly
sinuous posteriorly, with anterior insertion plate bearing 5
distinct slits which continue as shallow grooves leading to
anterior edge of tegmentum; tegmentum occupying approx-

Figs. 128-134. Choneplax lata (Guilding, 1829). Fig. 128. Whole specimen, 22.4 mm; Settlement Point, Grand Bahama; FSBC | 32547. Fig.
129. Juvenile, 6.5 mm; West End, Grand Bahama; FSBC | 32548. Fig. 130. Valve i ex 13.0 mm specimen; same lot as 129. Fig. 131. Valve
iv, Same specimen. Fig. 132. Valve viii, same specimen, dorsal view. Fig. 133. Same valve viii, lateral view. Fig. 134. Tegmental pustules,

valve iv, Same specimen (field width = 315 um).

LYONS: CARIBBEAN ACANTHOCHITONIDAE 109

imately 85% of valve length. Valves ii-vii elongate (Fig. 131),
strongly produced posteriorly to overhang following valves;
tegmentum elongate, pentagonal, widest behind middle, with
straight anterolateral margins; sutural laminae long, nearly
in line with plane of valves, curving anteromedially from
posterolateral corners of tegmentum, with subacute tips
separated anteriorly by deep, U-shaped sinus; single, shallow,
notch-like slits along lateral margins. Valve viii tegmentum pen-
tagonal (Fig. 132), widest anteromesially, produced posteriorly,
with mucro at posterodistal tip (Fig. 133); jugum absent;
sutural laminae extending tooth-like from anterolateral margins
of tegmentum; posterior insertion plate and slits absent.
Tegmental morphology of small specimens varies con-
siderably from that of larger specimens (Figs. 135-142). Valves
of very large specimens usually so eroded that posterior edges
are straight instead of pointed.

Jugum of valves ii-vii smooth, relatively narrow, little

138 142

Figs. 135-142. Choneplax lata (Guilding, 1829). Valves i-viii ex 6.7 mm
juvenile; West End, Grand Bahama; FSBC | 32548.

expanded anteriorly; jugum indistinct on valves of small
specimens. Tegmentum of all valves covered evenly with
coarse, spatulate pustules (Fig. 134) approximately 90 um
long, 50 um wide, generally flattened but with raised, central
dome and adapical macresthete, few or no micresthetes.

Girdle upper surface covered with small (100 »m),
densely packed, club-shaped spicules; anterior and sutural
tufts poorly developed, comprised of about 18-22 short (to 1.0
mm), stout, smooth, sharp-tipped, reddish brown or some-
times white spicules; marginal spicules 500 um long, smooth,
straight or slightly curved, white, rarely reddish brown; under-
side covered with small (100 um), straight, sharp-tipped, clear
spicules.

DISCUSSION: Choneplax lata is distinguished from all
species of Acanthochitona by lacking slits on the posterior
margin of valve viii (Figs. 143-145). Kaas (1972) and Ferreira
(1985) discussed uncertainty regarding the number of slits on
valve i and intermediate valves. The three specimens |
dissected (6.7-30.0 mm) each had 5 distinct slits on valve i,
not 3 as reported by Pilsbry (1893), and single, notch-like slits
on intermediate valves. Based on the 5-slitted valve i,
Choneplax is more similar to Acanthochitona than to Crypto-
plax, which has 3 slits; however, Choneplax shares the unsilit
tail valve with Cryptoplax.

Chiton strigatus Sowerby, 1840, has long been

144 145

Figs. 143-145. Choneplax lata (Guilding, 1829). Valves viii, ventral
views. Fig. 143. 6.7 mm specimen, same as Fig. 142 (specimen crack-
ed during handling). Fig. 144. 13.0 mm specimen, same as Fig. 132.
Fig. 145. Ex approximately 30.0 mm specimen (curled); West End,
Grand Bahama; FSBC | 32546.

110 AMER. MALAC. BULL. 6(1) (1988)

recognized as a later name for Choneplax lata. Status of Chiton
hastatus Sowerby, 1840, is less certain; most of the described
characters seem to indicate relationship to Choneplax, but
Carpenter (/n Pilsbry, 1893) examined the type specimen and
reported 2 slits in valve viii, indicating a species of
Acanthochitona.

Even though valve morphology changes considerably
with growth, Choneplax lata specimens of all sizes can be
recognized readily. Consequently, Kaas’ (1972) illustrations of
C. lata are perplexing. Drawings of a specimen from St. John,
Virgin Islands (Kaas figs. 108-112: ‘‘9 x 6.5 mm, curled’’) depict
a valve iv considerably wider than long, with short sutural
laminae, and a valve viii with a jugum and with lateral margins
of relatively short sutural laminae flush with those of the
tegmentum, which is posteriorly truncate. Although the unsilit
insertion plate seems identical to that of C. /ata, other il-
lustrated features differ markedly from valves iv and viii of the
6.7 and 13.0 mm specimens from Grand Bahama illustrated
here (see Figs. 131, 132, 138, 142, 143, 144). | did not illustrate
dorsal views of valves from larger specimens because they
inevitably were eroded. However, | did dissect a large speci-
men; most valves were posteriorly truncate but, except for
valve ti, the sutural laminae were relatively longer, not shorter,
than those of valves illustrated, and the tegmentum was
always longer than wide.

Kaas’ photograph (1972: pl. 2, fig. 4), reportedly of a
10.5 mm dried specimen of Choneplax lata from Spaanse
Water, Curagao, is difficult to interpret but does not much
resemble C. /ata. | did not examine any of the specimens Kaas
reported from the Virgin Islands, Tobago, Bonaire, or
Piscadera Baai and Spaanse Water, Curagao. However, | did
examine five uncatalogued RMNH lots of small specimens
(2.7-7.3 mm) labeled C. /ata from Aruba and Curagao, including
Piscadera Baai and Spaanse Water. Those lots all contained
specimens of Acanthochitona zebra, a species which
resembles C. /ata in the number, color, and shape of dorsal
tuft spicules and by the underhung insertion plate of valve
viii. Kaas also reported only small specimens (4-11 mm), and
characters he described on specimens from Bonaire and
Curagao could apply as well to A. zebra as to C. /ata. | am
not certain that specimens of both species were not mixed
in his account.

Ferreira (1985) ascribed greater morphological varia-
tion to small specimens of Choneplax lata than actually ex-
ists. Inexplicably, he decided that Acanthochitona andersoni,
A. balesae, and A. interfissa were juveniles of C. /ata. That
conclusion was incorrect, as demonstrated in preceding
treatments of those taxa. A simple proof, in addition to de-
scribed differences, is obtained by comparing valves viii. All
of the above Acanthochitona species, regardless of size, have
2 slits and an obvious jugum on valve viii, whereas even very
small (6.7 mm length) C. /ata lack any indications of posterior
slits or a jugum.

Ferreira’s confusion renders his distributional records
of Choneplax fata unreliable. Among IRCZM and CAS
specimens he identified, | found specimens of Acanthochitona
andersoni, A. balesae, and A. zebra as well as true C. /ata.
The illustrated specimen he tentatively labeled Choneplax cf.

lata from Barbados appears to be A. worsfoldi. Those
discrepancies are noted in the appropriate species accounts,
but many more lots must be re-examined before all of the
records can be corrected.

There seems to be no valid record of Choneplax lata
from Florida, perhaps because acceptable habitat does not
occur there. Specimens | collected at three locations in Grand
Bahama and Cuba lived along high energy rocky shores
washed by oceanic waves. Pilsbry (1893) described
specimens of C. /ata as vermiform, an apt descriptor consider-
ing their tendency to live in small round holes bored into large
limestone rocks.

Genus Cryptoconchus Burrow, 1815
Cryptoconchus floridanus (Dall, 1889)
Figs. 146-149

Notoplax floridanus Dall, 1889b: 416.

Acanthochites (Cryptoconchus) floridanus, Pilsbry, 1893: 37,
38, pl. 3, figs. 63, 64.

Cryptoconchus floridanus, Thiele, 1910: 110. Kaas, 1972: 34-36,
figs. 55-57, pl. 1, figs. 4, 5.

MATERIAL EXAMINED: FLORIDA: 2 spec., 10.8, 13.4 mm, patch
reef near Long Key Reef, Dry Tortugas, 1.5-2.5 m, 11-12 May 1979,
FSBC | 32074. —3 spec., 10.9-14.7 mm, Long Key Reef, Dry Tortugas,
intertidal, 11-12 May 1979, FSBC | 32075. —1 spec., 10.1 mm, Bird
Key Reef, Dry Tortugas, 0.5-1.0 m, 4 Oct 1979, FSBC | 32079. —1 spec.,
10.7 mm, Bird Key Harbor, Dry Tortugas, 2 m, 21 Aug 1981, FSBC
| 32487. —4 spec., 6.1-10.6 mm, Garden Key, Dry Tortugas, 1-2 m,
13 May 1979, FSBC | 32076. —1 spec., 11.3 mm, Garden Key, 1-2 m,
5 Oct 1979, FSBC | 32080. —3 spec., 7.3-12.8 mm, Key West, CAS
063314. —1 spec., 9.5 mm, West Summerland Key, 0-1 m, 27 Sept
1981, FSBC | 32082. —1 spec., 5.4 mm, Missouri Key, 0.5-1.0 m, 25
July 1987, FSBC | 32491. —4 spec., 9.2-14.6 mm, north side Vaca
Key, 1 Oct 1979, FSBC | 32077. —5 spec., 4.0-14.9 mm, northeast end
Vaca Key, 0-1.5 m, 4 Aug 1980, FSBC | 32081. —1 spec., 9.2 mm,
same location, 28 Sept 1981, FSBC | 32083. —5 spec., 7.2-11.1 mm,
Bonefish Key, CAS 063313. —1 spec., 14.1 mm, north side Grassy
Key, 0.5 m, 1 Oct 1979, FSBC | 32078. —2 spec., 5.0, 9.9 mm, east
end Grassy Key, 0-1 m, 18 Mar 1968, FSBC | 6395. —4 spec., all
curled, Burnt Point, Crawl Key, 2.5 m, July 1982, FSBC | 32488. —4
spec., all curled, same locality, 4 Aug 1982, FSBC | 32489. BAHAMAS:
1 spec., 13.2 mm, McLeanstown, east end Grand Bahama, 1-2 m,
24 May 1981, FSBC | 32486. —2 spec., 6.0, 13.0 mm, same locality,
27 Aug 1984, FSBC | 32084. —1 spec., curled, Georgetown, Great
Exuma, 21 June 1974, FSBC | 32526. TURKS AND CAICOS
ISLANDS: 1 spec., 13.6 mm, Providenciales, 0-2 m, 22 Sept 1986,
FSBC | 32490. PUERTO RICO: 1 spec., 14.0 mm, 2 km east of La
Parguera, 1m, 17 Aug 1985, FSBC | 32085. —1 spec., 13.0 mm, Cayo
Enrique, La Parguera, 1 m, 19 Aug 1985, FSBC | 32086.

DISTRIBUTION: Dry Tortugas and Florida Keys, Bahama
Islands to Puerto Rico, Cuba, Jamaica, and the Cayman
Islands, Aruba and Bonaire.

DESCRIPTION: Largest specimen 14.9 mm long, 8.9 mm wide
including girdle; valves nearly entirely covered by smooth,
black, brown, gray (rarely rose or yellow) girdle (Figs. 146, 147).
Narrow, white longitudinal bars evident in jugal region.
Exposed parts (jugum) smooth, that of valve i
semiovate, slightly wider than long; exposed jugal parts of

LYONS: CARIBBEAN ACANTHOCHITONIDAE 111

146 147

148 149

Figs. 146-149. Cryptoconchus floridanus (Dall, 1889). Fig. 146. Whole specimen, 12.4 mm; Vaca Key, Monroe County, Florida; FSBC | 32081.
Fig. 147. Whole specimen, 10.7 mm; same lot. Fig. 148. Jugum, valves iii-iv, same specimen as 147 (field width = 940 um). Fig. 149. Rudimen-
tary pustules bordering jugum; same specimen as 147 (field width = 175 um).

valves ii-vii narrow, straight-sided for about 60% of length,
thereafter expanded to truncate distal end, slightly elevated
at central posterior beaks; valve viii narrow, straight-sided, with
small, expanded, bulb-like terminus at low mucro. Tegmen-
tum virtually absent on valves, only occasionally represented
by few ovate, elongate pustules up to 80 nm long, 50 um wide
arranged parallel to jugal bars (Figs. 148, 149).

Girdle smooth, appearing granulose or warty under
magnification; 18 anterior and sutural dorsal pores situated
as in other Acanthochitonidae, bearing about 10 extremely
slender, fine-tipped spicules up to 100 um long; spicules at
peripheral margin sparse, short (40 »m), with blunt tips; under-
side densely covered with short (70-80 ym), sharp-tipped
spicules.

DISCUSSION: Pilsbry (1893) described the disarticulated
valves of Cryptoconchus floridanus as white, pink or purple;
the intermediate valves are rectangular, with a sinus before
and behind; there are 5 anterior slits on valve i, single slits
on the sides of valves ii-vii, and 2 posterior slits on valve viii.
Specimens examined herein, when viewed through the fleshy
underside, generally agreed with the standard 5-1-2 slit for-
mula. However, the largest specimen (FSBC | 32081) has 6
unevenly spaced slits on valve i. Tegmental pustules have not
been described for C. floridanus, but rudimentary pustules
sometimes do occur on valves where the girdle does not ex-
tend flush with the margin of the jugum.

The Florida range of Cryptoconchus floridanus has not
been extended since Dall’s (1889b) original description of
specimens from Cape Florida, Key Largo, Key West, and Dry
Tortugas. The species occurs throughout the Bahama Islands
and Greater Antilles, including Puerto Rico, Cuba (Jaume and

Sarasua, 1943), Jamaica (Humfrey, 1975), and the Cayman
Islands (Abbott, 1958). In the southern Caribbean, C.
floridanus has been reported from Aruba and Bonaire (Kaas,
1972). The species has not been reported in the western Carib-
bean from Mexico to Colombia.

DISCUSSION

More specimens must be examined before definitive
conclusions can be made on the composition and relation-
ships of the Acanthochitonidae of the Caribbean region. Of
the 14 recognized species, only Cryptoconchus floridanus has
not been involved in long-term or recent taxonomic confusion.
Thus, nearly all published records must be considered ques-
tionable, and the specimens upon which those records were
based must be re-examined. In addition, more collections of
Acanthochitonidae need to be made in the Lesser Antilles and
along the Caribbean coasts of Central and South America.
| examined far more material from Florida and the northern
Caribbean region than from the southern Caribbean. That im-
balance also occurs in published literature and probably will
be found in the unreported museum collections. To my
knowledge, there is no published record cf any polyplaco-
phoran from the area between Roatan, Honduras, and Limon,
Costa Rica, yet that area contains the vast, shallow Honduras-
Nicaragua shelf which exceeds in size the Bahama Banks.
Given those cautions, some observations on the Caribbean
Sea Acanthochitonidae seem warranted.

Occurrence of species may be limited by distributional
barriers, habitat availability, and environmental stress near
the northern boundary of the Caribbean region. Only Acan-
thochitona pygmaea occurs at Bermuda. In fact, only six of

112 AMER. MALAC. BULL. 6(1) (1988)

approximately fifty known species of shallow-water Caribbean
Polyplacophora occur at Bermuda (Jensen and Harasewych,
1986). The paucity of species at Bermuda probably is due
to long-term climate fluctuations and geographic isolation.

Seven species of Acanthochitonidae (Acanthochitona
anaersoni, A. balesae, A. hemphilli, A. pygmaea, A. roseo-
jugum, A. zebra, and Cryptoconchus floridanus) are known
from Florida, and it is unlikely that many more will be found
there. Most of the species are restricted to tropical en-
vironments of the Florida Keys and southeast coast and do
not occur in the more temperate environments of northeast
and west Florida. There are no endemic species. Previous
Florida records of A. astrigera and Choneplax lata are known
or suspected to be erroneous; | have collected both species
at various Caribbean locations, but | know of no similar
habitats where they could occur in Florida.

The northern Caribbean fauna, which extends from
Grand Bahama Island to Puerto Rico, the Virgin Islands, and
northern Netherlands Antilles (St. Eustatius and Saba Bank)
in the east and to Belize and Roatan in the west, is quite
diverse. Eleven species are known in that fauna, including
all of the Florida species plus Acanthochitona astrigera, A. line-
ata, A. worsfoldi, and Chonoplax lata. All eleven species have
been collected at Grand Bahama Island, and all except A.
worsfoldi have been collected at other northern Caribbean
sites. It is likely that intensive collecting will reveal similar
species richness at other northern Caribbean locations.

Only Acanthochitona andersoni, A. astrigera, and
Choneplax lata are known with certainty from the Lesser An-
tilles south of the northern Netherlands Antilles. However, Fer-
reira (1985) reported A. rhodea from Barbados, so it would
appear that a species of the A. hemphilli complex occurs there,
and Ferreira’s Barbados records of C. lata seem to be A.
worsfoldi.

The southern Netherlands Antilles (Aruba, Bonaire and
Curac¢ao) fauna is known to contain Acanthochitona ander-
soni, A. astrigera, A. balesae, the curiously restricted A.
bonairensis, A. rhodea, A. zebra, Choneplax lata, and Cryp-
toconchus floridanus. A total of eight species is indicated.

The fauna of southern Caribbean coastal areas is
poorest known. Along the entire expanse from Limon, Costa
Rica to Trinidad, | have seen only specimens of Acantho-
chitona andersoni, A. balesae, A. rhodea, A. venezuelana, and
a single specimen of an unknown Acanthochitona species
from Galeta Island, Panama.

There is little evident relationship between Brazilian
species of Acanthochitonidae and those of the Caribbean
fauna. Only three of the seven species of Acanthochitona re-
ported from Brazil were described from the Caribbean region,
and Brazilian records of each of those three species are ques-
tionable. Statements of the Brazilian occurrence of A.
spiculosa originally derived from E. A. Smith’s (1890) report
of A. astrigera at Fernando Noronha, but Righi (1971) also
reported A. spiculosa from off Sao Paulo in 25 m depth, far
deeper than the intertidal and shallow subtidal habitat other-
wise known for A. astrigera. A report of Brazilian specimens
of A. pygmaea by Righi (1971) was accompanied only by il-
lustrations of spicules and radula and was published when

the identity of that species was poorly understood; verified
specimens of A. pygmaea have been seen only from Saba
Bank northward to Bermuda. Brazilian records of A. hemphilli
are based on specimens reported from depths of 47-115 m
(Righi, 1971), whereas verified specimens have been seen on-
ly from Honduras and Puerto Rico northward to Florida and
from depths not greater than 18 m.

None of the four species of Acanthochitona originally
described from Brazil has been encountered in the Caribbean
fauna, and each has characters which distinguish it from any
Caribbean species. Acanthochitona brunoi Righi, 1971, has
a broad, strongly furrowed jugum bounded by only small
lateral tegmental areas whose anterolateral margins are con-
cave. The jugum of A. ciroi Righi, 1971, is very broad, occu-
pying more than half the total width of intermediate valves,
but is smooth, not furrowed, and valve i has fine, rib-like rows
of pustules (among other pustules) radiating toward the slits
from the posteromedial margin of the tegmentum. Pustules
of the tegmentum of A. minuta (Leloup, 1980) continue fully
formed over the entire jugum. The jugum of A. terezae Guerra
Junior, 1983, is similarly ill-defined and covered with pustules,
but the most distinctive features of that species occur on valve
viii, where the forward extension of the small, rudimentary
sutural laminae is far exceeded by that of the broad,
anterolaterally constricted tegmentum.

Several taxonomic groups are evident among the Car-
ibbean and eastern Pacific Ocean species of Acanthochitona.
Each group, or species complex, contains two Caribbean and
one eastern Pacific species as indicated by morphological
similarities. Closely related species complexes recognized
here include Acanthochitona hemphilli and A. rhodea (Carib-
bean) and A. ferreirai (eastern Pacific); A. pygmaea and A.
venezuelana (Caribbean) and A. avicula (eastern Pacific); and
A. astrigera and A. lineata (Caribbean) and A. hirudiniformis
(eastern Pacific). Watters (1981) proposed another species
complex containing Acanthochitona andersoni and A. balesae
(Caribbean) and A. arragonites (Carpenter, 1857) (eastern
Pacific). | have no study material of A. arragonites and so can-
not verify that relationship.

Relationships among the other Caribbean species of
Acanthochitona are less evident. Valve morphology of A.
roseojugum resembles that of species in the A. hemphilli com-
plex, and valves of A. worsfoldi resemble those of species
in the A. astrigera complex. However, girdle spicules of A.
roseojugum and A. worsfoldi hardly resemble spicules of
species in those complexes, so only remote relationships to
those species are proposed. A. bonairensis most resembles
the European A. fascicularis and does not much resemble
any other New World species.

The curious, underhanging posterior insertion plate
with two nearly vestigial slits, as well as the form, number,
and color of girdle spicules, suggest a relationship between
Acanthochitona zebra and Choneplax lata. However, their
resemblance probably represents convergence rather than
close phylogenetic relationship. Only single species of
Choneplax and Cryptoconchus, both Caribbean, are known
in the New World.

Distributional patterns of taxa in two of the Acantho-

LYONS: CARIBBEAN ACANTHOCHITONIDAE 113

chitona species complexes are known sufficiently to allow
speculation on their evolutionary history. A. rhodea and A.
venezuelana each occurs only along the southern Caribbean
coast, and each has a very similar cognate (A. ferreirai and
A. avicula) in the eastern Pacific region, as well as less similar
but still closely related congeners (A. hemphilli and A.
pygmaea) in the northern Caribbean. These distributional pat-
terns suggest at least two isolation-speciation events. In the
first event, A. pygmaea (and probably A. hemphilli) diverged
from the still-connected southern Caribbean-Panamic stocks
before or during the Pliocene, as evidenced by A. pygmaea
valves in Tertiary deposits. The valve reported as A. spiculosa
from the North Carolina Pliocene [Berry, 1940: 213, pl. 10 (not
pl. 12), figs. 5, 6] is not of A. pygmaea. However, Dall (1903),
who previously reported A. pygmaea as A. spiculosa, listed
both A. pygmaea and A. spiculosa in the Pliocene Caloosa-
hatchie beds of south Florida. The first isolation event left the
ancestors of A. pygmaea (and probably A. hemphilli) in the
northern Caloosahatchian fauna and left species resembling
A. avicula and A. rhodea in the southern Gatunian fauna (sen-
su Petuch, 1982). The known southern distributional limits of
A. hemphilli (Honduras) and A. pygmaea (Saba Bank) occur
precisely where Petuch (1982) identified areas of abrupt faunal
shift between the northern and southern Caribbean fauna.
Emergence of the Isthmus of Panama in the late Pliocene pro-
vided the barrier which resulted in later speciation among the
avicula-like and rhodea-like progenetors.

Speciation mechanisms in the Acanthochitona astri-
gera-lineata-hirudiniformis species complex are less evident.
The Caribbean A. lineata and eastern Pacific A. hirudinifor-
mis are most similar in valve morphology and thus seem to
have diverged most recently. To date, A. lineata is known on-
ly from the northern Caribbean, whereas A. astrigera occurs
in both the northern and southeastern Caribbean. Ferreira
(1985) reported A. astrigera from Caribbean Panama, but he
included three species (A. astrigera, A. lineata, and A. zebra)
within his concept of A. astrigera. Re-examination of his
Panama material might provide additional clues to the evolu-
tionary history of this species complex.

ACKNOWLEDGMENTS

Dr. Robert C. Bullock, University of Rhode Island, made
available to me many important specimens from his research col-
lection. Type-specimens and other material were provided by Piet
Kaas, Rijksmuseum van Natuurlijke Historie; Kathie Way, British
Museum (Natural History); and Dr. Robert Robertson, Academy of
Natural Sciences of Philadelphia. Dr. Terrence Gosliner provided
specimens from the California Academy of Sciences; Ms. Paula Mik-
kelsen provided specimens from the Indian River Coastal Zone
Museum; and Dr. Emily H. Vokes provided specimens from the Tulane
University Department of Geology. Dr. Pamela Hallock, University of
South Florida, St. Petersburg, provided specimens obtained by her
and her students while collecting foraminiferans in Florida and Puerto
Rico. Other specimens were provided from the private collections
of Margaret (Peggy) Williams, Sarasota, Florida; Robert Granda,
Panama City, Florida; Kevan Sunderland, Sunrise, Florida; Jack N.
Worsfold, Freeport, Grand Bahama Island; and Robert Quigley, also
of Freeport. My wife, Carol, worked with me to collect many of the

FDNR specimens upon which this study was based. Lana Tester and
Earnest Truby, both FDNR, produced the SEM photographs, and Sally
D. Kaicher, St. Petersburg, photographed many of the intact
specimens. James F. Quinn, Jr., and Thomas H. Perkins, both FDNR,
provided advice on nomenclature. All are thanked for their assistance.

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Date of manuscript acceptance: 23 October 1987

CHITONS (MOLLUSCA: POLYPLACOPHORA) FROM THE COASTS OF
OMAN AND THE ARABIAN GULF

PIET KAAS
RIJKSMUSEUM VAN NATUURLIJKE HISTORIE, P. O. BOX 9517,
2300 RA LEIDEN, THE NETHERLANDS

AND

RICHARD A. VAN BELLE
KONINKLIJK BELGISCH INSTITUUT VOOR NATUURWETENSCHAPPEN,
VAUTIERSTRAAT 29, 1040 BRUSSELS, BELGIUM

ABSTRACT

Twelve species of chitons are reported from the coasts of Oman and the Arabian Gulf. Mis-
identifications are corrected for five of the seven species previously reported from that area. New records
for the region include Lepidozona luzonica (Sowerby, 1842), Callistochiton adenensis Smith, 1891, Chiton
fosteri Bullock, 1972, Tonicia (Lucilina) sueziensis (Reeve, 1847), and Onithochiton erythraeus Thiele,
1910. Two new species, Acanthochitona woodwardi sp. nov., and Notoplax arabica sp. nov., are described.

The chiton fauna of the Arabian Gulf, the Gulf of Oman
and the Oman coast of the Arabian Sea has not been in-
vestigated thoroughly. Melvill and Standen (1901, 1906), re-
porting upon the mollusks of the Persian Gulf, Gulf of Oman
and Arabian Sea, did not mention any species of Polyplaco-
phora. Biggs (1958) reported Chiton lamyi Dupuis, 1917 (=
C. peregrinus Thiele, 1910) and C. (Acanthopleura) haddoni
from the Arabian Gulf, the latter from Hormuz Island, Iran,
at the entrance of the Gulf. Bosch and Bosch (1982: 145),
reported a single species, Acanthopleura haddon/ Winckworth,
1927 (= A. vaillantii de Rochebrune, 1882), a common rock-
dweller in the Red Sea and on the coasts of the northern In-
dian Ocean (except in the Arabian Gulf, where it is uncom-
mon). Those authors admitted that ‘‘there are several species
of chitons to be found in Oman, but most are small and pre-
sent problems in identification.’ Their specimens, collected
principally at the island of Masirah in the Arabian Sea, were
identified provisionally by Kathleen R. Smythe. Smythe (1982)
enumerated eight species of chitcns but, in glaring contrast
to the fine color photographs of gastropod and bivalve shells,
she illustrated the chitons with primitive line-drawings, by
which none are recognizable. Apart from several misidentifica-
tions, Smythe should be credited for establishing the occur-
rence of several well known and easily recognizable species
from the Arabian Gulf. Glayzer et al. (1984) listed five species
of chitons from Kuwait, including four listed previously by
Smythe. In the present study we establish the occurrence of
twelve species of chitons in littoral waters of the western

Arabian Gulf, the western Gulf of Oman, and the Oman coast
of the Arabian Sea.

HABITAT

A. J. Woodward provided the following descriptions of
the Qatar stations where he collected chitons, mostly by
snorkelling or scuba-diving. Ras Abruk (Fig. 1: no. 9) is a
sheltered bay on the end of a peninsula. The predominantly
limestone cliffs are ca. 10 m high, with raised fossil beds and
sandy beaches. Large boulders of limestone and aggregates
occur in the extreme shallows due to rock falls from the cliffs.
Fasht (= limestone and aggregate slabs with shells, pieces
of coral, etc.) occurs close to the shoreline and out to 30-40 m
in a broad broken band that is frequently exposed at low tide
and rarely covered by more than 30 cm of water. Beyond the
fasht band there is a small drop-off of mostly weed-covered
rocks, to a depth of about 2 m. Further offshore the substratum
is composed of fine sand for about 300 m, beyond which coral
and rock occur at a depth of 3-5 m. Chitons were always found
in the fasht band, where summer temperatures are extreme-
ly high (50+°C). Therefore, the water temperature is often
40+°C in the shallows. Salinity is similarly high (40+ ppt by
estimate).

Fuwairat (also spelled Fuwairet) (Fig. 1: no. 10) is a
coastal location with 30 m high limestone cliffs and small bays
at Jebel Fuwairat, about 1 km north of the village. Pebbly,
loose rocks, that occur at the extreme edge of the white sand

American Malacological Bulletin, Vol. 6(1) (1988):115-130

115

116 AMER. MALAC. BULL. 6(1) (1988)

4 SAWER KUWAIT

6 RAS AL JILA‘AT are

8 RAS DUKHAN aon
9 RAS ABRUK if

10 FUWAIRAT OATAR
11 AL WAKRAH

12 LAS HATTE 0

13 AS SHAAM UAE
14 QURM

15 AL BASTAN
16 RASSIER OMAN

17 HAQL
18 KURIA MURIYA |S

Goa”

pee |

Fig. 1. Map of Oman and the Arabian Gulf.

beach, are frequently exposed at low tide (maximum tidal
range ca. + 1.5m). Anarrow band of clean sand about 20 m
from shore can also be exposed during extreme low tides.
Further offshore the substratum is composed of loose rock
with some weed and algae cover, coral debris, small pieces
of live coral and fine sand. Seaward, small patches of live coral
occur on soft sand that becomes gravelly sand near coral
heads. An extensive coral reef is located ca. 200 m from shore
at a depth of 5 m. Chitons are found in the loose rock zone
about 30-75 m from shore, usually on undersides of rocks or
dead coral where the water depth rarely exceeds 1 m.
Temperature is very high in the shallows, though in winter it
drops below 11°C. This gives an annual temperature differen-
tial of ca. 30°C.

Wakrah is a small town south of Doha (Fig. 1: no. 11)
with very wide sandy beaches backed by limestone ridges.
In the extreme shallows near the beach broken fasht lies on
top of shelly gravel and sand. Hard packed, fine, white, ex-
posed sand bars are located about 50 m from the beach and
extend to ca. 200-300 m from the shoreline. These are ex-
posed at low tide. Following a slight drop-off into 1-1.5 m
depths, there first occurs a band of fine white sand, then loose
rock and dead coral covered by algae and weed. A second

fine white sand area occurs beyond the first and is followed
by another band of shell and coral debris and loose algae
and weed covered rocks that rise to ca. 30 cm in height. Here
chitons were occasionally found on the undersides of rocks.
Chitons were also found on shells of Pinna muricata L. The
chitons live on the parts of the shells which are deeply buried
in the sand. Chitons were never found on the broken fasht.
The temperature is high, ca. 2°C less than that at Ras Abruk;
the salinity is possibly higher.

Las Hatte (= Al Ashat), situated offshore from Umm
Said (Fig. 1: no. 12), consists of a group of four small limestone
islands with sandy beaches fringed by live coral about 75 m
from shore where a fairly steep drop-off occurs. Chitons
[= Lepidozona luzonica (Sowerby, 1842)] are found on dead
valves of arkshells (Arcidae) from about 10 m down to the
seabed at about 25-28 m. The substratum comprises a mix-
ture of silty black mud and sponges. Salinity is ca. 40-50 ppt
at the surface and increases with depth. The water
temperature in summer is lower than at other locations, rare-
ly exceeding 36°C; temperature in winter is ca. 12-15°C due
to greater water depth.

The following abbreviations are used throughout the
text: BG, Private collection of B. Glayzer; BMNH, British
Museum (Natural History), London; FH, Private collection of
F. Hinkle; KS, Private collection of K. Smythe; MCZ, Museum
of Comparative Zoology, Harvard University, Cambridge,
Massachusetts; MNHN, Muséum National d’Histoire
Naturelle, Paris; RMNH K, Private collection of P Kaas, now
in Rijksmuseum van Natuurlijke Historie, Leiden; VB, Private
collection of R. Van Belle; ZMHU, Zoologisches Museum an
der Humboldt Universitat, Berlin.

SYSTEMATIC ACCOUNTS
Class Polyplacophora
Order Neoloricata
Suborder Ischnochitonina
Family Ischnochitonidae Dall, 1889
Subfamily Ischnochitoninae
Genus /schochiton Gray, 1847

Type Species: Chiton textilis Gray, 1828 (by subsequent
designation, Gray, 1847).

Subgenus /schnochiton s.s.

Ischnochiton (I.) yerburyi (E. A. Smith, 1891)
Figs. 2-7

Chiton (Ischnochiton) yerburyi E. A. Smith, 1891: 420, pl. 33:
fig. 6.

Ischnochiton yerburyi, Pilsbry, 1892: 101, pl. 20: fig. 11. Nier-
strasz, 1905: 30. Thiele, 1910: 111, 113. Kaas, 1954: 5.
Leloup, 1960: 35, fig. 5. Biggs, 1973: 374. Leloup, 1980:
10. Smythe, 1982: 83, fig. 17. Ferreira, 1983: 251, figs.
1, 2. Glayzer et a/., 1984: 324. Kaas, 1986: 11, figs. 8,
8a, b (synonymy).

(?) Ischnochiton rufopunctatus Odhner, 1919: 21, pl. 3:
figs. 40, 41.

(?) Ischnochiton (Radsiella) delagoaensis Ashby, 1931: 40,
pl. 6: figs. 63-66.

Ischnochiton haersoltei Kaas, 1954: 5, figs. 7-9.

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF

117

Table 1. Distributional records of Polyplacophora in the Arabian Gulf and Oman. Species marked with an asterisk (*) also occur on the African

coast of the Indian Ocean.

Arabian Gulf Oman

Species Kuwait Bahrain Qatar U. A. E. Gulf of Oman Arabian Sea
*Ischnochiton yerburyi Smith, 1891 + + + = 4: we
I. winckworthi Leloup, 1936 + - + + = =
Lepidozona luzonica (Sowerby, 1842) - + ep + = =
Callistochiton adenensis Smith, 1891 = - - - = 7
Chiton peregrinus Thiele, 1910 + - + a + +
*C. fosteri Bullock, 1972 = = = = ae a
*C. (Rhyssoplax) affinis Issel, 1869 + - + i + =
*Acanthopleura vaillantii de Rochebrune, 1882 = (+)! - (+)! (+)! +
*Tonicia (Lucilina) sueziensis (Reeve, 1847) + + + - - +
*Onithochiton erythraeus Thiele, 1910 ~ - as = _ 4
Acanthochitona woodwardi sp. nov. + - + — = =
Notoplax (Notoplax) arabica sp. nov. + - + - = =

1Reported by Biggs (1958: 271) from Hormuz Id., Iran, at entrance of Arabian Gulf. Collected by Smythe (in litt. 3 June 1987) on the Trucial coast
of the Emirates, just inside the Gulf, at Khor Khaymal and Sharjah, and also at a point in Bahrain (!). Collected by Woodward at Dubai.

SYNTYPES: BMNH 1888.4.9.345.

MATERIAL EXAMINED. KUWAIT: 1 spec., 11.5 x 5.5. mm, Bide
Circle, under stones in tidepool, F. Hinkle leg., 12 June 1978, FH; —1
spec., ca. 7 mm long, id., 20 Sept 1979, FH; —3 spec., max. 15 x
8mm, id., 1 Aug 1981, FH; —2 spec., Kuwait Bay, on Pinna muricata,
intertidal, 19 Sept 1975, B. Glayzer leg., BG 1427; —Numerous valves,
Bahrain, in shell grit on beach, Nov 1971, F. van Nieulande don., VB
2667a. QATAR: 1 spec., 9 x 5 mm, Ras Abruk, under broken slabs
of fasht, intertidal, May 1982, A. Woodward leg., KS; —6 spec., max.
10.5 x 5.5 mm, Fuwairat, on rocks and dead coral, 0-1 m, June 1985,
A. Woodward leg., 4/KS, 2/RMNH K5105 (one disarticulated). OMAN:
3 spec., Gulf of Oman, Qurm, K. Smythe leg., 1979, KS; —2 spec.,
Arabian Sea, Masirah Id., Rassier, KS; —3 spec., Haql, K. Smythe
leg., KS.

TYPE LOCALITY: Aden.

DISTRIBUTION: Indo-Arabian coasts from Karachi, Pakistan,
to Aden in Yemen, and in the Red Sea to the Gulf of Aqaba,
Israel; African coast from Somalia to Zanzibar (many of these
records are unconfirmed).

DESCRIPTION: This taxon was adequately described and il-
lustrated by E. A. Smith (1891: 420, pl. 33 fig. 6) except for
details of girdle armature and radula which follow (see also
Figs. 2-5). Dorsal girdle scales (Fig. 6) broadly rounded,
moderately curved, ca. 100 x 80 um, with 12-15 elevated,
slightly converging riblets separated by somewhat narrower,
rather deep grooves.

Central tooth of radula (Fig. 7) narrow, abruptly widen-
ing distally to umbrella-like blade; first lateral teeth as long
as central tooth, narrow, with inwardly curved, roundish blade;
major lateral teeth with bidentate head, inner cusp much
stronger than outer one, shaft with a short, trunk-like appen-
dix just under and before head; spatulate uncinal with blunt-
ly pointed, outwardly incised cusp.

DISCUSSION: Ferreira (1983: 251) combined all Indian Ocean
species of /Ischnochiton with ‘‘reticulate, thimble-like
sculpture.’ Whether he was correct in synonymizing /schno-
chiton sansibarensis Thiele, 1910, /. delagoaensis Ashby, 1931,

|. kilburni Kaas, 1979, from Mozambique, and /. rufopunctatus
Odhner, 1919, from Madagascar, with /. yerburyi cannot be
decided here. Close reexamination of the types could reveal
a complex of sibling species, rather than one variable species.
As far as we can ascertain, /. haersoltei Kaas, 1954, from
Manora Island, Karachi, does not differ from Gulf specimens
of /. yerburyi.

Ischnochiton (I.) winckworthi Leloup, 1936
Figs. 8-15

Ischnochiton winckworthi Leloup, 1936: 51, figs. 1-9, 1949: 1,
figs. 1, 2, 3A, 4-7, pl. 1; 1952: 15. Rajagopal and Subba
Rao, 1974: 404, 409. Smythe, 1982: 83, fig. 16.

Ischnochiton ranjhai Kaas, 1954: 8, figs. 10-14.

SYNTYPES: BMNH.

MATERIAL EXAMINED: KUWAIT: 1 spec., 7.5 x 4 mm, Bide Cir-
cle, under stones in tidepool, F. Hinkle leg., 20 Sept 1979, FH; —2
spec., max. 5 x 3.5 mm, id., 1 Aug 1981, FH; —2 spec., max. 7 x
4 mm, id., 10 Sept 1983, FH; —1 valve + girdle, Sawer, 1974, K.
Smythe leg., KS; —1 spec., Kuwait Bay, 14 Feb 1975, B. Glayzer leg.,
KS. QATAR: 2 spec., Ras Dukhan, 15 Apr 1978, K. Smythe leg., KS.
—9 spec., max. 10 x 5.5 mm, Ras Abruk, under broken slabs of fasht,
intertidal, May 1982, A. Woodward leg., 7/KS, 2/RMNH K5097. —2
spec., Ras Abruk, 2-3 Nov 1978, A. Partridge leg., KS. —3 spec., max.
10 x 5.5 mm, Fuwairat, on rocks and dead coral, 0-1 m, June 1985,
A. Woodward leg., KS. U. A. E.: 2 spec., 3.2 and 2.6 mm long, Abu
Dhabi, K. Smythe leg., KS.

TYPE LOCALITY: Sri Lanka, near Trincomali, Dutch Bay.

DISTRIBUTION: Locally common along the shores of
Malaysia, Andaman Islands, Burma, Sri Lanka, Pakistan,
Kuwait, Qatar, U. A. E.; intertidal.

DESCRIPTION: Animals small, ca. 10 mm long, width ca. 2/3
length, largest specimen recorded 15 x 9.5 mm (Leloup, 1936:
51), oval, moderately raised (dorsal elevation 0.35-0.41),
carinated, side slopes straight to slightly convex, valves not
beaked. Color of tegmentum variable, beige, olivaceous, dark

118 AMER. MALAC. BULL.

Figs. 2-7. Ischnochiton yerburyi Smith (specimens from Fuwairat, Qatar, Apr 1985, A. Woodward leg. in coll. Smythe, RMNH K5105). Fig. 2.
Valve IV, dorsal view, 3.7 mm wide. Fig. 3. Valve VIII, dorsal view, 3.7 mm wide. Fig. 4. Camera lucida sketch of valve IV, rostral view, 5.5 mm
wide. Fig. 5. Lateral view of valve VIII, 2.4 mm wide. Fig. 6. Dorsal girdle scale. Fig. 7. Central, first lateral, major lateral and spatulate uncinal

radula teeth.

greyish green, with roughly symmetrical blotches of dirty white
on central part of valves. Many specimens with 2-3 dark spots
at posterior margin of valves, some specimens uniformly
roseate, more exceptionally, white or brownish.

Head valve (Fig. 8) semicircular, front slope straight,
posterior margin widely V-shaped, weakly notched medially.
Intermediate valves (Figs. 9, 10, 13) broadly rectangular, front
and hind margins nearly straight, parallel-sided, apices hardly
or not indicated, side margins rounded, lateral areas little
raised but neatly marked. Tail valve (Figs. 11, 12) somewhat
less than semicircular, mucro not prominent, slightly anterior,
posterior slope concave.

Tegmentum granulose, sculpture often obsolete in
younger specimens, variable in older ones. In most commonly
occurring form, head valve of adult specimen sculptured with
36-40 radiating, somewhat irregular, granulose riblets, becom-
ing obsolete toward apex, growth lines hardly or not indicated,
lateral areas of intermediate valves with 4-5 similar radiating
riblets, some bifurcating near outer margin, central areas, and
antemucronal area of tail valve, with weak, fine, longitudinal
riblets, 10-15 per side, becoming obsolete toward the finely
quincuncially granulose jugal area, postmucronal area of tail
valve sculptured like head valve.

Articulamentum whitish to light roseate, tegmental color
visible, apophyses thin, sharp, moderately wide, evenly
arched, jugal sinus straight, ca. 1/5 width of valve, insertion

plates short, slit formula 8-11/ 1/ 9-10, slit rays finely indicated,
teeth sharp, smooth, eaves solid.

Girdle moderately wide with alternating bands of
yellowish and greyish green, dorsally covered with strongly
bent, imbricating scales, ca. 150 x 120 wm; top rounded,
ornamented with ca. 10 strong ribs wider than interstices (Fig.
14). Margin with fringe of short, white, torpedo-shaped
spicules. Ventral side of girdle paved with radiating rows of
elongate rectangular, smooth scales, 67 x 20 um. Radula (Fig.
15) with narrow central tooth bearing a roundish, upwardly
curled blade; first laterals equally narrow, ending in inwardly
curved, hook-shaped blade; major laterals with strong, sharply
pointed main cusp and short minor denticle on outside. Gills
holobranchial, abanal, 18 ctenidia per side in 74 mm
specimen.

Genus Lepidozona Pilsbry, 1892

Type Species: Chiton mertensii von Middendorff, 1847 (by
original designation).
Subgenus Lepidozona s.s.
Lepidozona (L.) luzonica (Sowerby, 1842)
Figs. 16-23

Chiton luzonicus Sowerby, 1842: 104. Reeve, 1847: pl. 25:
sp. and fig. 167. Van Belle, 1982: 473.

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 119

15 \

Figs. 8-15. /schnochiton winckworthi Leloup [Figs. 8-10: paratype of /schnochiton ranjhai Kaas, 1954 (H. Heyn, del.), RMNH K3422; Figs. 11-15,
specimen from Ras Abruk, Qatar, May 1982, A. Woodward leg. in coll. Smythe, RMNH K5097]. Fig. 8. Valve I, dorsal view, 3.7 mm wide. Fig.
9. Valve III, dorsal view, 3.5 mm wide. Fig. 10. Camera lucida sketch of valve VIII, rostral view, 3.8 mm wide. Fig. 11. Dorsal view of valve
Vill, 4.7 mm wide. Fig. 12. Lateral view of valve VIII, 2.7 mm wide. Fig. 13. Rostral view of valve IV, 5.3 mm wide. Fig. 14. Dorsal girdle scale.
Fig. 15. Central, first lateral, major lateral and spatulate uncinal girdle teeth.

Ischnochiton (Lepidozona) luzonicus Pilsbry, 1893: pl. 38,
figs. 31-32; 1894: 85.

Ischnochiton luzonicus Nierstrasz, 1905: 34. Hidalgo, 1905:
271; Faustino, 1928: 123.

Callistochiton finschi Thiele, 1910: 86, pl. 8: figs. 57-60; 1911:
402. Ashby, 1923: 236. Iredale and Hull, 1925: 354. Fer-
reira, 1974: 163; 1978: 39.

Solivaga finschi |redale and Hull, 1925: 355, pl. 40: figs. 14-16.
Cotton, 1964: 55.

Lorica (Solivaga) finschi Thiele, 1929: 18.

Lepidozona luzonica Kaas and Van Belle, 1987: 245, fig. 111,
map 52.

non /Ischnochiton (Lepidozona) luzonicus Ang, 1967: 401,
pl. 5: figs. 1-5 (= Chiton sp.).

LECTOTYPE: BMNH 1979. 175/1 (by subsequent designation,
Kaas and Van Belle, 1987).

MATERIAL EXAMINED: BAHRAIN: 1 valve, in shell grit on beach,
Nov. 1971, F. van Nieulande don., VB 2975a. QATAR: 2 spec., Fuwairat,
June 1985, A. Woodward leg., 1/KS 1/RMNH K5100. —4 spec., Las
Hatte, on dead shells, 10-20 m, 26 July 1985, A. Woodward leg., 2/KS,
1/RMNH K5099, 1/VB 2975b (disarticulated); —7 valves (mounted on
slide) Fuwairat or Las Hatte, June/July 1985, A. Woodward leg., KS.
U. A. E.: 1 spec. (in alcohol), Abu Dhabi, K. Smythe leg., 4.2 mm
long, KS.

TYPE LOCALITY: Philippines, province Albay, Isle of Luzon,
Sorsogon, 27 m.

DISTRIBUTION: Eastern coast of Sumatra (Java Sea),
Singapore (as Callistochiton finschi), Bahrain, Qatar and
U. A. E.

DESCRIPTION: Animal small, lectotype (Fig. 16) 9.2 x 5.8 mm,
largest specimen 12 x 7 mm (Iredale and Hull, 1925: 355, as
Solivaga finschi), oval, moderately elevated (dorsal elevation

120 AMER. MALAC. BULL. 6(1) (1988)

Figs. 16-23. Lepidozona luzonica (Sowerby) [Fig. 16, lectotype: Figs.
17-23, paralectotypes (BMNH 1979.175)]. Fig. 16. Whole specimen,
dorsal view, 5.8 mm wide. Fig. 17. Right half of valve Ill, dorsal view,
4.6 mm wide. Fig. 18. Camera lucida sketch of valve Ill, rostral view,
7.3 mm wide. Fig. 19. Valve Ill, dorsal view, 7.3 mm wide. Fig. 20.
Valve Il, dorsal view, 6.9 mm wide. Fig. 21. Dorsal girdle scales. Fig.
22. Central and first lateral radula teeth. Fig. 23. Heads of major lateral
teeth.

0.36-0.39) carinated, side slopes straight, valves not beaked.
Color of tegmentum yellowish to greenish with, on central
areas, few longitudinal streaks of darker tone, or buff, sparsely
spotted with bluish green.

Head valve semicircular, front slope somewhat con-
cave, hind margin widely V-shaped, deeply notched in mid-
dle, tegmentum sculptured with low, radial, often bifurcating,
granulose riblets, 40-50 in number along outer margin, becom-
ing obsolete toward apex. Intermediate valves (Figs. 17-20)
broadly rectangular, front and hind margins straight, parallel-
sided, side margins rounded, apices inconspicuous, lateral
areas little raised, 5-6 riblets, up to 7-9 by splitting, central
areas with 12-16 longitudinal, granulose ridges per side, ridges
close-set and little pronounced on jugal areas, gradually more
widely spaced and elevated toward side margins, interspaces
finely, densely, but irregularly, transversely grooved. Tail valve
subsemicircular, almost as wide as head valve, mucro at
anterior third of valve, not prominent, postmucronal area rather
flat, sculptured like head valve, ca. 32 riblets along outer
margin, antemucronal area sculptured like central areas.

Articulamentum glossy white, apophyses very wide,
short, rounded, connected across shallow sinus by short,

slightly concave, laminated jugal plate, weakly notched at
sides, slit formula 11-14/ 1/ 10-13, slits inequidistant, slit rays
indicated, teeth short, weakly grooved on outside, eaves nar-
row, solid.

Girdle buff-colored, sometimes banded with bluish
green, dorsally covered with obliquely implanted, slightly bent,
more or less rectangular scales, with 12-16 obsolete ribs, up
to 125 um long, 188 um wide in mid-girdle, smaller toward the
outer margin (Fig. 21).

Central tooth of radula (Fig. 22) narrow at base,
gradually widening to strong, rounded blade, first lateral tooth
about as long as central one, slender, with somewhat distorted
blade, major lateral (Fig. 23) with a tricuspid head, denticles
sharply pointed, central one longer than others.

DISCUSSION: The present specimens undoubtedly are con-
specific with Lepidozona luzonica, differing only in a less pro-
nounced sculpture; radula and girdle armature are exactly like
specimens of L. /uzonica from elsewhere. Specimens from
the Arabian Gulf extend the known range of L. luzonica con-
siderably to the west and establish the presence of Lepidozona
in the northwestern Indian Ocean.

Subfamily Callistoplacinae Pilsbry, 1893
Genus Callistochiton Carpenter in MS; Dall, 1879
Type Species: Callistochiton palmulatus Carpenter in MS
(by monotypy, Dall, 1879).

Callistochiton adenensis (E. A. Smith, 1891)
Figs. 24-27

Chiton (Callistochiton) adenensis E. A. Smith, 1891: 421,
pl. 33: fig. 7.

Callistochiton adenensis Pilsbry, 1893: 276, pl. 59: fig. 45.
Nierstrasz, 1905: 41. Sykes, 1907: 31. Thiele, 1910: 84,
pl. 8: figs. 49-51. Ashby, 1923: 233. Leloup, 1952: 30;
1953: 1, fig. 1. Kaas, 1979: 861. Ferreira, 1979: 463.
Zeidler and Gowlett, 1986: 114.

Lepidopleurus rochebruni Jousseaume, 1893: 102. Nier-
strasz, 1905: 10; 1906: 145, 157.

HOLOTYPE: BMNH.

MATERIAL EXAMINED: OMAN: 2 spec., max. 24 x 12 mm, Al
Bastan or Masirah Id., Mar 1984, D. Bosch leg., 1/KS, 1/RMNH K5101.
—1 spec., 16 mm (curled), Arabian Sea, Masirah Id., |. 1984, D. Bosch
leg., KS; —1 spec., id., between Haql and Rassier, K. Smythe leg.,
KS. —1 spec., 18 mm, Rassier, K. Smythe leg., KS. —2 spec., 18.5.
18 mm long, (disarticulated), Rassier, 9 Feb 1982, K. Smythe leg.,
1/KS, 1/VB 2976a.

TYPE LOCALITY: Aden.

DISTRIBUTION: Gulf of Aden; Arabian coast of Oman;
possibly Gulf of Oman.

DESCRIPTION: Girdle densely covered with strongly im-
bricating, wide, short, oval, curved scales, with more than
twenty elevated riblets, narrow, latticed interstices, ca. 140 x
50 um; marginal scales small and narrow, bluntly conical, 25
x 50 um, with ca. 6 ribs; ventral side of girdle covered with
transverse rows of rectangular scales, ca. 60 x 15 um (Figs.
24-26).

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 121

ci.

Figs. 24-27. Callistochiton adenensis (Smith) (specimen from Oman,
Masirah Id. or Al Bastan, Mar 1984, D. Bosch leg. in coll. Smythe,
RMNH K5101). Fig. 24. Valves 1-3 in situ, 12 mm wide. Fig. 25. Ven-
tral girdle scales. Fig. 26. Dorsal girdle scales. Fig. 27. Central, first
lateral and major lateral radula teeth.

Central tooth of radula (Fig. 27) somewhat pinched in
middle, with semi-oval, rather narrow blade; first laterals
somewhat S-shaped, embracing central tooth, with broad ex-
terior wing in basal part and small rounded blade; major
laterals with bicuspid head, denticles stout, sharply pointed,
interior one slightly longer, shaft with short, curved appendix
at inside of head; spatulate uncinals with narrow, rounded cut-
ting edge.

Short, poorly illustrated original description of this
species was amplified by Thiele (1910) who produced good
figures of the valves, and by Leloup (1953), who also figured
the girdle elements.

Family Chitonidae Rafinesque, 1815
Subfamily Chitoninae
Genus Chiton Linnaeus, 1758
Type Species: Chiton tuberculatus Linnaeus, 1758 (by subse-
quent designation, Dall, 1879).
Subgenus Chiton s.s.
Chiton (C.) peregrinus Thiele, 1910
Figs. 28-30

Chiton (Clathropleura) peregrinus Thiele, 1910: 90, pl. 9:
figs. 23-27.

Chiton lamyi Dupuis, 1917: 538. Biggs, 1958: 271. Smythe,
1982: 82, fig. 15. Glayzer et al., 1984: 324.

Chiton lamyi var. reticulatus Dupuis, 1918: 532.

Chiton wallacei Winckworth, 1927: 206, pl. 29: figs. 5-8.

Chiton iatricus Winckworth, 1930: 78, pl. 8b. Smythe,
1982: 82.

Chiton iatricus var. winckworthi Kaas, 1954: 2.

Chiton peregrinus Bullock, 1972: 238, pl. 44: figs. 1, 2, 10
(bibliography and synonymy). Ferreira, 1983: 268.
Zeidler and Gowlett, 1986: 113.

SYNTYPES: ZMHU.

MATERIAL EXAMINED: KUWAIT: 4 juv. spec., Falaika Id., Al Zor,
on rocks, intertidal zone, 10 Nov 1975, B. Glayzer leg., BG 1428 (as

Chiton lamyi). QATAR: 3 spec., Dasa, K. Smythe leg., KS. U. A. E.:
—1 partly disarticulated spec., As Shaam, K. Smythe leg., KS. OMAN:
6 spec., max. 37 x 22 mm, Al Bastan or Masirah ld., Mar 1984, D.
Bosch leg., KS. —3 spec., Gulf of Oman: Qurm, 1979, K. Smythe leg.,
KS. —2 spec., max. 30 x 17 mm, Muscat, Mar 1969, D. Bosch leg.,
VB 2651a. —2 spec. + partly disarticulated + 1 disarticulated red
spec. + 6 valves, Arabian Sea, Masirah ld., 12 Jan 1984, D. Bosch
leg., KS. —3 spec. + 8 valves, id., Rassier, K. Smythe leg., KS. —23
spec., max. 28 x 20 mm (slightly curled), between Rassier and Haq],
K. Smythe leg., KS. —3 spec., Haql, K. Smythe leg., KS.

TYPE LOCALITY: S Africa, ? Algoa Bay (in error = Aden,
fide Bullock, 1972).

DISTRIBUTION: Widely distributed in the northwestern Indian
Ocean from the north coast of western India to the Arabian
Gulf and westward to the entrance of the Red Sea; intertidal
in rocky areas.

DESCRIPTION: Specimens large, up to 7 cm long, greater
than 4 cm wide. Shells, older animals, typically strongly erod-
ed; young specimens with two thread-like radial riblets on
lateral areas, one accompanying diagonal mark, another at
short distance from posterior margin (Fig. 28). Tegmentum
always granulate, granules on central areas arranged in
somewhat wavy series perpendicular to diagonal lines, con-
verging toward jugum. Color mostly greyish green, sometimes
with black markings, valves of disarticulated specimen
(Masirah Id., Oman) reddish all over along with articula-
mentum.

Girdle paved with strong, large, imbricating scales, with

lozenge-shaped base, strongly bent, smooth on outside if not
eroded (Fig. 29); scales ca. 0.75 mm wide, slightly less high,
bluntly pointed at top. Central tooth of radula very narrow,
sagittate, first laterals broad at base, narrowing distally, without
blade; major laterals with simple, oval head without cusp (Fig.
30).
DISCUSSION: This species was well described by several
authors who, owing to intraspecific variation and state of
preservation, created different names for it. The complicated
synonymy was Clearly established by Bullock (1972). It is by
far the most common chiton on the Indo-Arabian coasts.

Chiton (C.) fosteri Bullock, 1972
Figs. 31-33

Chiton fosteri Bullock, 1972: 245, pl. 44: figs. 6-9. Kaas,
1979: 862.

HOLOTYPE: MCZ 279166.

MATERIAL EXAMINED: OMAN: 1 spec., 40.5 x 21 mm, Arabian
Sea, Masirah Id., Haql, K. Smythe leg., KS.

TYPE LOCALITY: Madagascar, Ile Ste Marie, Ankoalamare.

DISTRIBUTION: Madagascar, Mozambique, the Comoro
Archipelago, Zanzibar and Kenya; locally common.

DESCRIPTION: Single specimen bluish green with faint
zebra-pattern of brownish concentric lines (Fig. 31). Slit for-
mula 9/1/16.

122 AMER. MALAC. BULL. 6(1) (1988)

ann 1S
1ooum

\ ; y

‘/ 30

lew /
‘ Wes 1, op ————
“ 100
7 | 7 seoum

Figs. 28-30. Chiton peregrinus Thiele (Fig. 28: specimen from Manora Id., Karachi, Pakistan, 15 Feb 1953, S. M. H. Bilgrani leg., RMNH K4705;
Figs. 29-30: specimen from Oman, Masirah Id., between Haq! and Rassier, K. Smythe coll.). Fig. 28. Valve V, dorsal view, 8.9 mm wide. Fig.
29. Dorsal girdle scales, left dorsal, right ventral view. Fig. 30. Central, first lateral and major lateral radula teeth.

Dorsal girdle scales (Fig. 32) spindle-shaped, base
elongate, lozenge-shaped, dorsal surface strongly convex, ap-
parently smooth. Under high magnification scales appear fine-
ly punctate-lineate toward base, minutely bubbled around top.
Scales on mid-girdle measure ca. 680 x 300 xm. Radula (Fig.
33) central tooth almost linear, with narrow, sagittal blade;
major laterals closely packed, with oval head, edge of free
margin sharp.

DISCUSSION: This species was well described by Bullock
(1972). Additional observations were added by Kaas (1979).

Subgenus Rhyssoplax Thiele, 1893

Type Species: Chiton janeirensis Gray, 1828, sensu Thiele,
1893 (= Chiton affinis \ssel, 1869) (by subsequent designa-
tion, L.C.Z.N., 1971).

Chiton (Rhyssoplax) affinis |ssel, 1869
Figs. 34-40

Chiton affinis Issel, 1869: 234. Beu et a/., 1969: 184. Yaron,
1973: 15. Sabelli, 1974: 75. Fischer, 1978: 43.
Lepidopleurus bottae de Rochebrune, 1882: 192. Ferreira,
1983: 270, fig. 24.

Callistochiton heterodon savignyi Pilsbry, 1893: 277, pl. 60:
fig. 16. Ferreira, 1983: 270.

Chiton olivaceus var. affinis, Leloup, 1952: 27, fig. 11, pl. 4:
fig. 4. (bibliography and synonymy); 1960: 36. Sabelli
and Spada, 1970: 6.

Callistochiton barnardi, Smythe, 1982: 81, fig. 14 (non Ashby,
1931). Glayzer et a/., 1984: 324.

Rhyssoplax affinis, Ferreira, 1983: 268, fig. 22.

LECTOTYPE: MNHN (by subsequent designation, Ferreira,
1983).

MATERIAL EXAMINED: KUWAIT: 2 spec., max. 12 x 6 mm, Bide
Circle, under stones in tidepool, F. Hinkle leg., 20 Sept 1979, FH; —3
spec., max. 11.5 x 5 mm, id., 1 Aug 1981, FH; —2 spec., max. 10
x 5.5. mm, id., 10 Apr 1983, FH; —6 spec. (as Callistochiton barnardi),
Kuwait Bay, on underside of rocks, intertidal zone, 19 Sept 1975, B.
Glayzer leg., 5/BG 1426, 1/KS (disarticulated). QATAR: 10 spec., max.
15 x 8 mm, Ras Abruk, under broken slabs of fasht, intertidal, May
1982, A. Woodward leg., 6/KS, 2/RMNH K5104, 2/VB 2768b; —4 spec.
(one heavily damaged), Ras Abruk, 3 Nov 1978, A. Partridge leg.,
KS. —13 spec., max. 14 x 6.5 mm, Fuwairat, on rocks and dead cor-
al, 0-1 m, June 1985, A. Woodward leg., 12/KS, 1/RMNH K5103; .2
spec., Al Wakrah, K. Smythe leg., KS. OMAN: 2 spec., Gulf of Oman,
Qurm, 1979, K. Smythe leg., KS.

TYPE LOCALITY: Gulf of Suez.

DISTRIBUTION: Gulf of Suez, Red Sea and Somalia
(southernmost record Sar Uanle), Arabian Gulf, the Gulf of
Oman; intertidal to shallow subtidal.

DESCRIPTION: Dorsal girdle scales regularly imbricating, im-
planted in cuticula of girdle by diamond-shaped base, strongly
curved dorsally, round-topped, ornamented with ca. 8 broad
flat, weakly convergent ribs separated by narrow grooves, ca.
285 x 140 um (Figs. 34-39).

Radula (Fig. 40) with narrow central tooth, blade nar-
rowly U-shaped; first laterals broad at base, in middle with
wing-like procession on inner sides, abruptly narrowing distal-
ly, ending bluntly rounded without blade; major laterals with
broad, oval head, free margin sharply edged; on the inside
of it the shaft bears a slender, trunk-like appendix.

DISCUSSION: The quite extensive original description (Issel,
1869) has been supplemented by several authors. Leloup
(1952) produced detailed figures of the girdle elements. Yaron
(1973) demonstrated the consistent morphological differences

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 123

fe
100umM

Figs. 31-33. Chiton fosteri Bullock (specimen from Oman, Masirah
Id., Haql, K. Smythe leg. and coll.). Fig. 31. Whole specimen, dor-
sal view, 27.6 mm wide. Fig. 32. Dorsal girdle scales, above ventral
view, below dorsal view. Fig. 33. First and major radula teeth.

between Chiton affinis and the related Mediterranean Sea
species C. (R.) olivaceus Spengler, 1797. Ferreira (1983)
described the radula.

Subfamily Acanthopleurinae Dall, 1889
Genus Acanthopleura Guilding, 1829
Type Species: Chiton spinosus Bruguiére, 1792 (by subse-
quent designation, Gray, 1847).
Acanthopleura vaillantii de Rochebrune, 1882

Chiton testudo Spengler, 1797: 78 (nom. nud.).

Acantopleura (sic !) vaillantii de Rochebrune, 1882: 192.
Pilsbry, 1894: 97. Nierstrasz, 1906: 514. Winckworth,
1927: 206. Ferreira, 1983: 278; 1986: 226, 231, fig. 17.

Acanthopleura sp. (?) Haddon, 1886: 24.

Acanthopleura haddoni Winckworth, 1927: 206, pl. 28: figs. 1-4.
Leloup, 1937: 172, figs. 17-19; 1960: 38. Pearse, 1978:
95, fig. 2. Leloup, 1980: 6. Bosch and Bosch, 1982: 145,
fig. Smythe, 1982: 82. Ferreira, 1983: 278; 1986: 226,
227.

Chiton (Acanthopleura) haddoni, Biggs, 1958: 271; 1969: 201.

LECTOTYPE: MNHN (by subsequent designation, Ferreira,
1986).

TYPE LOCALITY: Suez Canal.

DISTRIBUTION: Red Sea, Yemen, Oman, Arabian Gulf at
Jumeira (near Dubay), U. A. E., Khor Khaymah (S of As
Shaam), U. A. E., Sharjah (N of Dubay), U. A. E., near Hor-
muz Id. and the opposite coast of Iran, also ‘‘at a point in
Bahrain’’ (K. Smythe, in litt. 3 June 1987), on rocky shores
and in rock pools.

DESCRIPTION: Animal large, to 75 mm long, width about 2/3

Figs. 34-40. Chiton (Rhyssoplax) affinis |Issel. (specimens from
Fuwairat, Qatar, June 1985. A. Woodward leg. in coll. Smythe, RMNH
K5102). Fig. 34. Whole specimen, dorsal view, 6.7 mm wide. Fig.
35. Camera lucida sketch of valve IV, rostral view, 3.5 mm wide. Fig.
36. Valve IV, dorsal view, 3.5 mm wide. Fig. 37. Valve VIII, dorsal view,
2.75 mm wide. Fig. 38. Camera luciaa sketch of valve VIII, lateral
view, 1.94 mm wide. Fig. 39. Dorsal girdle scales. Fig. 40. Central,
first lateral and major lateral radula teeth.

124 AMER. MALAC. BULL. 6(1) (1988)

the length, broadly oval, moderately raised, back almost
rounded, side slopes slightly convex, valves more or less
beaked, generally strongly eroded. Tegmentum dark reddish
to blackish brown, some specimens with traces of longitudinal
bands of lighter color on jugum.

Head valve nearly semicircular, front slope convex,
posterior margin concave in central part, convex toward sides.
Intermediate valves broadly rectangular to widely V-shaped,
side margins decidedly rounded, apices indicated, blunt,
lateral areas little or not raised, hardly marked. Tail valve less
than semicircular, crescentic in some specimens, as wide as
head valve, mucro somewhat raised, postmedian.

Tegmental sculpture, often indistinguishable on ac-
count of erosion and incrustation, consists of small, roundish
tubercles arranged in irregular, more or less concentrical rows.
Extra-pigmentary eyes very small, abundantly distributed on
end valves and more than half of lateral areas of intermediate
valves.

Articulamentum glossy, dark brown in central part of
valves, light greyish brown toward sides, apophyses large,
rounded, somewhat obliquely directed, connected across
sinus by short, concave jugal plate, insertion plates short, slit
formula 10/1/9-10, slits narrow, inequidistant, slit rays not in-
dicated, teeth finely but deeply grooved on dorsal side, pec-
tinate, those of tail valve slightly directed anteriorly.

Girdle wide, dark brownish, or whitish with irregular
dark brown bands, densely clothed dorsally with large, coarse,
blunt-pointed, calcareous spines of different forms and sizes,
interspersed with small, slender spicules. Marginal spicules
more or less cylindrical, almost as long as dorsal spines, blunt-
topped, with some wide, longitudinal ribs. Girdle paved ven-
trally with radiating rows very small, thick scales, slightly
longer than wide, squarish at base, distally tapering to blunt
top, ornamented with 4-5 strong, longitudinal ribs.

Central tooth of radula slenderly elongate, with broad,
strongly convex blade, first lateral tooth irregularly rectangular,
major lateral with large unicuspid head, denticle abruptly
pointed. Gills holobranchial, abanal.

DISCUSSION: This large, easily recognizable species, was
well described by several authors including Haddon (1886),
Winckworth (1927) and Leloup (1937). Winckworth produced
good figures of the complete animal and the loose valves,
and Leloup gave detailed figures of the girdle elements.
“Chiton punctatus L.’’ of Spengler (1797:76) is probably a
synonym. It was based on animals from the Red Sea, mostly
desicated and disarticulated specimens that have lost their
girdle armature, leaving deep pits in the cuticula (hence: punc-
tatus!). At the end of his description Spengler wrote (p. 78):
“‘The Arabian Society sent it from the Red Sea together with
other products of this sea. Due to the similarity of the valves
it might be called Chiton testudo’’ (translated from Danish).
Although A. vaillantii is the only representative of Acantho-
pleura in the Red Sea and Spengler’s specimens undoubtedly
belong to this genus, there is no certainty about their true iden-
tity, so the name C. testudo is to be regarded a nomen dubium,
leaving A. vaillantii the oldest available valid name.

In the opinion of Ferreira (1986), Acanthopleura vaillantii

should be regarded as another of the many synonyms of A.
gemmata (Blainville, 1825). Winckworth (1927), however, clear-
ly showed his A. haddoni (=vaillantii) to be different from A.
spiniger (Sowerby, 1840) [=A. gemmata (Blainville)] in several
respects; ‘‘in haddoni the valves are broader, the diagonal
ribs almost obsolete in adult and young specimens, the
sculpture is finer abd closer and is uniform over the central
and lateral areas, the tail valve is more rounded; the inser-
tion plates and sutural laminae are differently proportioned...’

Leloup (1937: 174) wrote: ‘‘The characteristics of the
girdle and of the tegmentum as far as the aesthetes are con-
cerned allow us to differentiate A. haddoni from A. spiniger’’
(Sowerby, 1840). On the one side the girdle of spiniger dor-
sally bears an underground of uniform, small, brown spicules,
which haddoni does not show; the thick spines are fairly
regularly equal in spiniger, whereas they are very irregular
in shape and of inequal dimensions in haddoni; the scales
of the ventral side are relatively longer in spiniger. On the other
side the aesthetes show the same general aspect, but they
are more globulous in haddoni, especially in the lateral areas”’
(translated from French). We agree with the arguments of
Winckworth and Leloup and retain A. vaillantii as a valid
species.

Subfamily Toniciinae Pilsbry, 1893
Genus Tonicia Gray, 1847
Type Species: Chiton elegans Frembly, 1827 (non de Blain-
ville, 1825) (= Chiton chilensis Frembly (1827) (by subsequent
designation, Gray, 1847).
Subgenus Lucilina Dall, 1882
Type Species: Chiton confossus Gould, 1846 (= Chiton
lamellosus Quoy and Gaimard, 1835) (by subsequent designa-
tion, Pilsbry, 1893).
Tonicia (Lucilina) sueziensis (Reeve, 1847)
Figs. 41-44

Chiton sueziensis Reeve, 1847: pl. 20, sp. and fig. 134.

Tonicia ptygmata de Rochebrune, 1883: 33. Ferreira, 1983:
274, fig. 28.

Tonicia sueziensis (sic!), Leloup, 1960: 40, figs. 6, 8, pl. 1, fig. 1
(bibliography and synonymy); 1973: 9, 18; 1980: 12.

Tonicia sueziensis, Kaas, 1979: 871. Ferreira, 1983: 271, figs.
25-27; Kaas, 1986: 18.

LECTOTYPE: BMNH 1951.2.7.7 (by subsequent designation,
Ferreira, 1983).

MATERIAL EXAMINED: KUWAIT: 2 spec., max. 12 x 7 mm, Bide
Circle, under stones in tidepool, F. Hinkle leg., 12 June 1978, FH. —3
spec., max. 9.5 x 6.5 mm, id., 1 Aug 1981, FH; —5 spec., max. 13
x 7.5 mm, id., 10 Apr 1983, FH. BAHRAIN: 1 valve, in shell grit on
beach, Nov 1971, F. van Nieulande don., VB 2610a. QATAR: 2 spec.,
max 9 x 6 mm (slightly curled), Fuwairat, on rocks and dead coral,
0-1 m, June 1985, A. Woodward leg., 1/KS, 1/RMNH K 5102. —1 spec.,
22.5 x 85 mm, AL Wakrah, on loose rocks covered with algae and
weed, 0-1.5 m, June 1984, A. Woodward leg., KS.

DISTRIBUTION: Gulf of Suez, Red Sea, coasts of Somalia,
Seychelles Is and Coetivy Id.; Kuwait, Bahrain and Qatar; in-
tertidal to shallow subtidal.

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 125

Figs. 41-44. Tonicia (Lucilina) sueziensis (Reeve) (specimen from
Fuwairat, Qatar, June 1985, A. Woodward leg. in coll. Smythe, RMNH
K5098). Fig. 41. Right half of valves IV and V in situ, 4.25 mm wide.
Fig. 42. Dorsal girdle spicules. Fig. 43. Ventral girdle scales. Fig.
44. Central, first and major lateral radula teeth.

TYPE LOCALITY: Egypt, Suez.

DESCRIPTION: Tonicia (Lucilina) sueziensis was adequately
described by several authors, particularly Leloup (1960), who
produced detailed figures of the girdle elements, and Ferreira
(1983), who gave good figures of the lectotype and an accurate
description of the radula.

Girdle covered dorsally with extremely minute, bullet-
shaped spicules, 36 x 20 um, top with 4-5 short riblets on visi-
ble half; ventral scales (Fig. 43) arranged in lateral rows, rec-
tangular, base slightly concave, top rounded, 25 x 19 um (Figs.
41-43).

Central tooth of radula (Fig. 44) small, very narrow, 68
x 7 um, with sharply pointed blade; first laterals broad, base
bluntly pointed, pinched in middle, gradually widening distaad,
with outwardly directed extension; major laterals with
tetracuspid head, denticles short, bluntly rounded, shaft with
trunk-like appendix just under and beneath head, directed
inward.

DISCUSSION: Ferreira (1983: 274) wrongly synonymized
Tonicia (Lucilina) carnosa Kaas, 1979, from Mozambique, the

Comoro Archipelago, and Madagascar, with the present
species. 7. (L.) carnosa differs considerably in color and in
having much weaker sculpture with far fewer longitudinal
grooves on central areas of intermediate valves.

Genus Onithochiton Gray, 1847
Type Species: Chiton undulatus Quoy and Gaimard, 1835. non
Olfers, 1818; Wood, 1828 (= Onithochiton neglectus de
Rochebrune, 1881 (by subsequent designation, Gray, 1847).

Onithochiton erythraeus Thiele, 1910
Figs. 45-50

Onithochiton erythraeus Thiele, 1910: 98, pl. 10, figs. 53-55.
Leloup, 1941: 13; 1960: 42, 45, 47. Glynn, 1970: 17.
Kaas, 1979: 872. Ferreira, 1983: 276-277.

Onithochiton lyelli forma erythraeus, Pearse, 1978: 93, 95,
fig. 3.

HOLOTYPE: ZMHU.

MATERIAL EXAMINED: OMAN: 1 spec., 16 mm, Arabian Sea,
Masirah Id., 12 Jan 1984, D. Bosch leg., KS. —1 spec., length 21 mm,
(slightly curled), id., Rassier, 9 Feb 1982, K. Smythe leg., KS. —2
spec., max. width 12.5 mm (both curled), between Rassier and Haq],
K. Smythe leg., 1/KS 1/RMNH K5098. —1 spec., length 13 mm
(curled), Haql, K. Smythe leg., KS.

TYPE LOCALITY: Erythraea, El Tor.

DISTRIBUTION: Gulf of Suez, Red Sea, and Arabian Sea
coast of Oman; intertidal.

DESCRIPTION: Girdle densely clothed dorsally with tiny,
bluntly pointed scales, ca. 23 x 10 um, top with 5-7 riblets (Figs.
45-49). Marginal spicules (Fig. 50) smooth, cylindrical, bluntly
pointed, ca. 90 x 22 um; ventral scales very small, shorter
than wide, ca. 20 x 15 um.

Central tooth of radula (Figs. 46, 47) about twice as long
as wide, widest in anterior part, with extremely narrow, abrupt-
ly pointed blade and median, raised riblet in anterior half, base
equally pointed; first laterals twice length of central teeth, trun-
cate at base, with central, short, sharp thorn, gradually nar-
rowing anteriorly, ending in narrow, rounded blade; major
laterals with tetracuspid head, denticles short, blunt, shaft with
short, funnel-shaped appendix just under and anteriorly of
head; spatulate uncinal teeth with elongate triangular blade.

DISCUSSION: Though it has not been studied thoroughly
before, Onithochiton erythraeus has been compared with
several related Onithochiton species. Leloup (1941) conclud-
ed it was synonymous with O. maillardi (Deshayes, 1863) from
Mauritius. Later, he (Leloup, 1960) considered both of these
species, as well as O. quercinus (Gould, 1846) from New South
Wales, O. literatus (Krauss, 1848) from South Africa, O.
wahibergi (Krauss, 1848) from the Cape of Good Hope, O.
rugulosus Angas, 1867 from New South Wales, and O. schol-
vieni Thiele, 1910 from New South Wales, to be junior
synonyms of O. /lyelli (Sowerby, 1832). Ferreira (1983)
synonymized O. wahibergi, O. maillardi and O. erythraeus with
O. literatus. Kaas (1979) expressed some doubts as to the con-

126 AMER. MALAC. BULL. 6(1) (1988)

Figs. 45-50. Onithochiton erythraeus Thiele. Fig. 45. Valves VI-VIII
in situ, dorsal view, 10.3 mm wide. Fig. 46. Central and first lateral
radula teeth. Fig. 47. Major lateral and spatulate uncinal teeth. Fig.
48. Dorsal girdle scales from mid-girdle. Fig. 49. Same, near outer
margin. Fig. 50. Marginal spicules.

clusions of Leloup (1960). Pending a thorough study of the
type material and good specimens from the different localities,
we prefer to treat the matter conservatively and consider O.
erythraeus a valid species, especially after we were able to
compare the Oman specimens with several lots of O. literatus
from Isipingo, Natal, and Inhaca Id., Lourengo Marques,
Mozambique, which proved to be quite differently sculptured.

Suborder Acanthochitonina
Family Acanthochitonidae Pilsbry, 1893
Subfamily Acanthochitoninae
Genus Acanthochitona Gray, 1821
Type Species: Chiton fascicularis Linnaeus, 1767 (by monotypy).

Acanthochitona woodwardi Kaas and Van Belle, sp. nov.
Figs. 51-60

TYPE MATERIAL: HOLOTYPE: 6.7 x 4.0 mm, Qatar, Dasa, 15 Nov
1978, K. Smythe leg., BM(NH) 1987032. PARATYPES: QATAR: 17
spec., max. 9.0 x 4.7 mm, collected with holotype, 13/KS, 2/RMNH
K5095 (one disarticulated, figured here), 2/VB 2968b. —2 spec.
(curled, one on matchstick) Ras Abruk, 3 Nov 1978, A. Partridge leg.,
KS. —1 spec. (disarticulated), Ras Abruk, under broken slabs of fasht,
intertidal, May 1982, A. Woodward leg., KS. —3 spec., Fuwairat, on
rocks and dead coral, 0-1 m, June 1985, A. Woodward leg., 2/KS
1/RMNH K5096. —2 spec. (one juvenile), Al Wakrah, K. Smythe leg.,
KS. KUWAIT: 2 spec., Al Bide, on rocks in intertidal zone, 29 Jan 1975,
B. Glayzer leg., 1/BG 1425 (as Chiton sp.), 1/KS (disarticulated, on
slide). —2 spec., 5.5 x 3.0 mm (damaged) and 5.0 x 2.5 mm (disar-
ticulated), Bide Circle, under stones in tidepool, MTL, F. Hinkle leg.,
12 June 1978, former, FH, latter, VB 2968a. —1 spec., 8.5 x 4.0 mm,
id., 1 Aug 1981, FH.

DISTRIBUTION: Kuwait and Qatar; intertidal.

DIAGNOSIS: Animal small, holotype 6.7 x 4 mm, length of
largest specimen 9 mm, width about half length, elongate
oval, rather flat (dorsal elevation 0.20-0.24), back subcarinate,
side slopes straight to slightly convex, head and intermediate
valves decidedly beaked. Tegmentum mostly whitish to light
beige, speckled or flecked with dark greyish green, some
specimens with light brownish or reddish brown, more or less
triangular blotch on jugum of valve II, another specimen red-
dish, shading into roseate to whitish on apical areas, holotype
blackish brown, with jugum and girdle whitish. Tegmental
sculpture of flat, roundish to oval, neatly separated granules,
jugal areas not raised, weakly ribbed longitudinally. Girdle
finely spiculose, little encroaching at sutures. Major lateral
radula tooth tricuspid.

DESCRIPTION: Head valve (Fig. 51) semicircular, front slope
somewhat convex, anterior margin vaguely waved, posterior
margin beaked, tegmentum sculptured with neatly separated,
flat, roundish to oval, quincuncially arranged granules, larger
toward outer margin, smaller, becoming obsolete toward apex,
no growth lines. Intermediate valves (Figs. 52-53) twice as
wide as long, front margin straight to slightly convex at both
sides of concave jugal part, hind margin concave at both sides
of strongly protruding apex, jugal area narrowly wedge-
shaped, not raised, sculptured with ca. 5 weak, flat,
longitudinal ribs separated by very fine grooves, lateral areas
not marked but slightly raised with regard to pleural areas,
sculpture of latero-pleural areas similar to that of head valve
but granules larger, more widely spaced, less regularly ar-
ranged. Tail valve (Figs. 54, 55) slightly oval transversely,
mucro prominent, pointed, somewhat behind centre, posterior
slope strongly concave, tegmentum sculptured like latero-
pleural areas of intermediate valves.

Articulamentum whitish, tegmental color slightly visi-
ble through, intermediate valves with transverse callus in cen-

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 127

55

100 UM

Figs. 51-60. Acanthochitona woodwardi sp. nov. Fig. 51. Valve |, dorsal
view, 3.33 mm wide. Fig. 52. Camera lucida sketch of valve IV, rostral
view, 4.44 mm wide. Fig. 53. Valve IV, dorsal view, 4.44 mm wide.
Fig. 54. Valve VIII, dorsal view, 3.38 mm wide. Fig. 55. Camera lucida
sketch of valve VIII, lateral view, 1.95 mm. Fig. 56. Dorsal girdle
spicules. Fig. 57. Small and large spicule from sutural tuft. Fig. 58.
Ventral spicules. Fig. 59. Central and first lateral radula teeth. Fig.
60. Blade of major lateral tooth. (Figs. 56-60, scale bar = 100 um.)

tral part, apophyses rounded, sharp, smooth, jugal sinus
about 1/5 valve width, weakly concave, insertion plates rather
short, slit formula 5/ 1/ 2 (figured specimen with only 4 slits
in valve l), slits shallow, slit rays hardly or not indicated, teeth
sharp, smooth to very finely striate, eaves solid.

Girdle densely covered dorsally with small, straight to
slightly bent, abrupty pointed, smooth spicules, ca. 60 x 10
um (Fig. 56), sutural tufts relatively short, composed of
straight, slender, sharply pointed, smooth spicules of different
sizes, varying from 180 x 8 um to 400 x 30 um (Fig. 57). Ven-
tral side of girdle paved with close set, radiating rows of sharp-
ly pointed, smooth spicules, up to 100 x 20 um (Fig. 58).
Marginal spicules similar to ventral ones.

Central tooth of radula (Fig. 59) elongate tulip-shaped,
with thin blade, first lateral tooth somewhat shorter, slender,
slightly widening distally, anterolateral corner with sharply
pointed outward lobe, no blade, major lateral with tricuspid
head, denticles pointed, central one longer than others (Fig.
60).

Gills merobranchial, abanal, 11 ctenidia per side.

DISCUSSION: This species cannot be attributed to any known
species in the Indian Ocean. Acanthochitona mahensis Winck-
worth, 1927, from Mahé, India, and A. ashbyi Leloup, 1937,
from the Indian Ocean (?), possibly a synonym of the former,
differ in their greater size, more close packed, coarser
granules, finely ribbed jugal areas, relatively wider tail valves
with convex postmucronal slope, and longer, coarser, long-
itudinally striate marginal girdle spicules. In A. curvisetosus
Leloup, 1960, from the Red Sea, the granules are smaller and
more rounded, the jugal areas relatively wider and orna-
mented with ca. 15 longitudinal striations, and, contrary to
those of A. woodwardi, the ventral girdle spicules are smaller
than the dorsal ones. A. limbata Kaas, 1986, from Madagascar,
differs in the form of the valves, the drop-shaped granules,
the much broader jugal areas, and the form of the major lateral
radula tooth.

ETYMOLOGY: This species is named after Mr. A. J.
Woodward.

Genus Notoplax H. Adams, 1861
Type Species: Cryptoplax (Notoplax) speciosa H. Adams, 1861
(by monotypy).
Subgenus Notoplax s.s.
Notoplax (N.) arabica Kaas and Van Belle, sp. nov.
Figs. 61-72

Schizochiton jousseaumei, Smythe, 1982: 84, fig. 18 (non
Dupuis, 1917). Glayzer et a/., 1984: 324.

TYPE MATERIAL: HOLOTYPE, 11.4 x 5.9 mm, Kuwait Bay, Kuwait,
on rocks and dead shells, intertidal, 14 Feb 1975, B. Glayzer leg.,
BMNH 1987031. PARATYPES: 2 spec., 9.9x 5mm and 8.3 x 3.9mm,
collected with holotype, BG 1195. —7 valves (disarticulated), collected
with holotype, RMNH K5106. —1 spec., 11.0 x 6.0 mm (disarticulated),
Fuwairat, Qatar, on rocks and dead coral, 0-1 m, June 1985, A. Wood-
ward leg., KS.

DISTRIBUTION: Kuwait, Qatar; intertidal.

DIAGNOSIS: Animal small, 10-11 mm long, width about half
length, rather flat, side slopes slightly convex, valves little
beaked. Tegmentum uniformly light ochraceous, greyish or
light orange, coarsely sculptured with large, strongly elevated,
radially directed, elongate pustules, jugal areas wedge-
shaped, raised, back rounded, neatly separated from the ad-
jacent lateropleural areas by deep, wide grooves. Girdle finely
spiculose, deeply encroaching between valves. Major lateral
radula tooth tricuspid.

DESCRIPTION: Head valve (Fig. 61) nearly semicircular, front
slope weakly convex, anterior margin with five more or less
distinct waves, posterior margin widely V-shaped, minutely
notched in middle, no trace of radial ribs. Intermediate valves
(Fig. 62) broadly triangular, front margin slightly concave at
both sides of strong, forwardly produced, convex jugal part,
hind margin weakly beaked, somewhat sinuose at sides,
apices sharply pointed, lateral areas indiscernible. Tail valve
(Figs. 63, 64) very small, transversely oval, mucro prominent,
not elevated, slightly postmedian, posterior slope deeply
concave.

128 AMER. MALAC. BULL. 6(1) (1988)

100 4M

“

‘mete

64 70 71

Figs. 61-72. Notoplax arabica sp. nov. Figs. 61-63. Valves | (4.0 mm
wide) V (5.4 mm wide) and VIII (4.0 mm wide) respectively, dorsal
view. Fig. 64. Valve VIII, ventral view (4.0 mm wide). Fig. 65. Heads
of major lateral teeth. Fig. 66. Spatulate uncinal tooth. Fig. 67. Cen-
tral and first lateral radula teeth. Fig. 68. Dorsal girdle spicules. Fig.
69. Marginal spicule. Fig. 70. Spicule from sutural tuft. Fig. 77.
Spicule from girdle bridge. Fig. 72. Ventral spicules (Figs. 65-67, scale
bar = 100 um; Figs. 68-72, scale bar = 10 um).

Tegmentum microscopically granulose, end valves and
lateropleural areas of intermediate valves sculptured with
large, strongly raised, convex, widely spaced, irregularly oval
to decidedly elongate, radially oriented pustules, those near
valve margins overhanging or projecting past valve, on some
valves pustules vaguely arranged in irregular, longitudinal
rows, jugal areas raised, smooth to naked eye, ornamented
with extremely fine, longitudinal, beaded riblets, accompanied
by shallow, longitudinal excavation on both sides.

Articulamentum slightly translucent, tegmental color
shining through, apophyses strongly forwardly produced,
rounded, jugal sinus moderately to strongly convex, insertion
plates long, slit formula 5/ 1/ 6, slits shallow, those of tail valve
inequidistant, slit rays hardly or not indicated, teeth of head
and intermediate valves long, sharp, weakly striate dorsally,
those of tail valve short, blunt, strongly striate.

Girdle dorsally covered with small, straight to slightly
bent, sharply pointed, faintly longitudinally striate spicules,
56-62 «m long, 8-10 um thick (Fig. 68), on girdle bridges in-
terspersed with long, slender, straight, smooth spicules, 110
x 8 wm (Fig. 71), sutural tufts composed of stout, straight,
sharply pointed spicules, 100 x 14 um, weakly longitudinally
striate on distal half (Fig. 70). Marginal spicules (Fig. 69) small,
decidedly obese, blunt-pointed, finely longitudinally striate,
52 x 14 um. Girdle paved ventrally with very small, slender,
straight spicules, 40 x 3 um (Fig. 72).

Central tooth of radula (Fig. 67) tulip-shaped, with
straight blade, first lateral tooth somewhat shorter, narrowly
aliform, without blade, major lateral with tricuspid head, den-
ticles pointed, central one longer than others (Fig. 65),
spatulate uncinal tooth bent, smooth, distal end rounded.

DISCUSSION: By its peculiar sculpture of large, elongate,
convex, widely spaced pustules, N. arabica differs markedly
from all known Notoplax spp. in the Indian Ocean, its closest
relatives being N. elegans Leloup, 1981, from Madagascar,
which has a greater number of close set, subcircular, con-
cave granules, N. alisonae (Winckworth, MS; Kaas, 1976), from
Sri Lanka, which has a much greater number of tear-shaped,
flat to slightly concave granules, and N. coarctata (Sowerby,
1841), from the Philippines, in which the tegmentum of in-
termediate valves is flask-shaped.

ETYMOLOGY: The name of this species reflects its presence
in the Arabian Gulf.

DISCUSSION

The most striking phenomenon among the present
material is the discontinuous distribution of Lepidozona
luzonica, hitherto known only from Luzon, Philippines, the
Java Sea and Singapore. Its occurrence in the Arabian Gulf
remains inexplicable except for transport resulting from
human intervention via navigation. This is the case with
Chaetopleura angulata (Spengler, 1797) and Acanthochitona
fascicularis (Linnaeus, 1767). Another striking fact is the ab-
sence of Chiton huluensis (Smith, 1903), also discontinuously
distributed, covering a vast area from the Tasman Sea through
the Torres Straits, the Moluccas, the Timor Sea, the Maldive
Islands, Sri Lanka, the western coast of Madagascar, the coast
of Mozambique, the Red Sea and through the Suez Canal
to the Mediterranean coast of Israel. It should be remembered,
however, that the bulk of material we studied was collected
in intertidal or shallow subtidal areas, except for some
specimens collected in 15-20 m depths by SCUBA.

Ferreira (1983) concluded that “‘at least as far as chiton
faunas are concerned, the tropical western Indian Ocean con-
stitutes a definite zoogeographic province, which includes the
Red Sea, the East African coast southward to Natal, and the
adjacent islands eastward to Mauritius (60°E).”’ Undoubtedly
the chiton fauna of the western Indian Ocean is far richer in
species than is that of the Indo-Arabian side. However, 50%
of the species found in the Gulf and on the Oman coast also
are found on the African coast, and all but one also occur
in the Red Sea. On the other hand, Callistochiton adenensis

KAAS AND VAN BELLE: CHITONS OF ARABIAN GULF 129

occurs on the Oman coast as well as in the Red Sea but has
not been found in Somalia nor south of there. So we can con-
clude that the Red Sea chiton fauna is composed both of
African and Indo-Arabian species. Nevertheless, a comparison
of the faunas of both sides of the Indian Ocean leads to some
preliminary conclusions. The genus Chiton s.s. is represented
by two species on both sides: C. peregrinus in the east, C.
salihafui Bullock, 1972, in the west, and C. fosteri on both sides.
Two other Indo-Arabian chiton species reach their northern
limit in the Gulf, /schnochiton winckworthi and Lepidozona
luzonica. Acanthopleura vaillantii appears to be the only
representative of that genus in the east, whereas it is accom-
panied by A. brevispinosa (Sowerby, 1840) on the African
coast. Two species of Cryptoplax have been reported from the
African coast, of which C. sykesi Thiele, 1909, also is found
in the Red Sea; none are found on the Indo-Arabian side.

Because of the limited number of chiton species oc-
curring in the northern tropical Indian Ocean, the distribu-
tion data reviewed here do not allow zoogeographic provinces
to be established.

ACKNOWLEDGMENTS

We express our gratitude to all persons who provided
specimens and collection data, without which this paper could not
have been written. We render special thanks to Mrs. Kathleen Smythe
of Bognor Regis, West-Sussex, U.K., who generously lent all the
chitons she collected in and outside the Arabian Gulf, together with
quite a lot collected by Mr. Anthony Woodward, now at Abu Dhabi,
U.A.E., and by Dr. Donald Bosch, then at Oman, who has done much
collecting in the Arabian Sea and the Gulf of Oman. Mr. Woodward
kindly provided ample information on his collecting sites in Qatar.
Finally we thankfully acknowledge Mrs. Barbara Glayzer of London,
who kindly lent her Kuwait specimens, and Mr. F. Hinkle of Kuwait,
who sent many fine chiton specimens to the junior author (VB) for
identification.

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Date of manuscript acceptance: 22 October 1987

SENSE ORGANS IN THE GIRDLE OF CHITON OLIVACEUS
(MOLLUSCA: POLYPLACOPHORA)

FRANZ PETER FISCHER, BRIGITTE EISENSAMER, CHRISTINA MILTZ
AND INGRID SINGER
INSTITUT FUR ZOOLOGIE, TECHNISCHE UNIVERSITAT MUNCHEN
LICHTENBERGSTRASSE 4, D 8046 GARCHING,
FEDERAL REPUBLIC OF GERMANY

ABSTRACT

A general scheme for the girdle sense organs in the Polyplacophora is put forward: a sensory
papilla, inserted in the girdle epithelium, consists of a varying number of secretory cells, one ciliary
cell and one spicule cell. The spicule cell is connected with an organic cup or shaft. In or on top of
this structure there is a calcareous element (spicule, scale or small tip). The ciliary cell invaginates
into the spicule cell. Around this invagination the cytoplasm of the spicule cell contains a dense net-
work of microfilaments and a large number of mitochondria.

Modifications of this scheme are found in Chiton olivaceus. Behavioral experiments demonstrate

that the girdle sense organs are mechanoreceptors.

The presence of sense organs (aesthetes and shell
eyes) in the shell valves of chitons has been known since the
work of Moseley (1884). The occurrence of sensory structures
in the girdle that surrounds the valves has only recently been
demonstrated (Haas and Kriesten, 1975; Fischer et a/., 1980;
Leise and Cloney, 1982; Leise, 1986). Previously, the hard
structures of the girdle were considered as armament and
ornamentation by most authors. However, Blumrich (1891) and
Plate (1898, 1902) suggested that some girdle formations could
be sensory. The ventral and dorsal surface differ in arrange-
ment and form of these structures, differences which are
species-specific. In addition, many species have different
spines along the girdle margin. In several families, hair-like
structures are also produced. A survey of the different forms
is given in Hyman (1967). Hyman also suggests that hairs
could be modified shafts of spicules.

The ultrastructure of the girdle is still poorly Known.
With the exception of Lepidochitona cinereus L. (Haas and
Kriesten, 1975), the species that have been studied, Acan-
thochitona fascicularis L. (Fischer et al., 1980) and Mopalia
muscosa Gould (Leise and Cloney, 1982; Leise, 1986), have
a highly specialized girdle.

This study concerned Chiton olivaceus Spengler,
the most common chiton in the Adriatic Sea; it is domi-
nant in the tidal and low subtidal region (Leloup and
Volz, 1938). The girdle ornamentation has been described
by Blumrich (1891) and no obvious specializations exist.
We examined the ultrastructure of both scales and hairs

in order to reveal the basic structure of the girdle sense
organs.

MATERIAL AND METHODS

One to three year old individuals of Chiton olivaceus
from the tidal zone of the coast of northern Yugoslavia were
used in this study. For transmission electron microscopy, parts
of the girdle were fixed in 5% glutaraldehyde in phosphate
buffer (pH 7.4) for two hours. They were then decalcified in
3% EDTA in phosphate buffer overnight following postfixation
in 2% osmium tetroxide for two hours. All this was done at
3°C. After dehydration (ethanol, propylene oxide) the
specimens were embedded in Durcupan and ultrathin sec-
tions cut with a Reichert ultramicrotome. The sections were
stained with uranyl acetate and lead citrate using the stan-
dard methods of Reynolds (1963) and studied in a Jeol elec-
tron microscope.

For scanning electron microscopy, some specimens,
after complete dehydration in a graded series of ethanol, were
critical point dried (Balzers CPD 020) from liquid carbon diox-
ide. Specimens were then coated with a 300 A layer of gold
and viewed in a Jeol 25S SEM.

In order to qualitatively test the reactions of the animals
to touching of single spines or other formations of the girdle,
a glass microelectrode filled with 3M KCL was connected via
a preamplifier to an audiomonitor. Because the sound fre-
quency changes due to a change in resistance of the

American Malacological Bulletin, Vol. 6(1) (1988):131-139

131

132 AMER. MALAC. BULL. 6(1) (1988)

Fig. 1. Schematic cross section through the girdle (a, articulamen-
tum; ae, aesthetes in the tegmental shell layer; c, cuticle; cl, clap-
per; ct, connective tissue; ds, dorsal scales; h, hair; ms, marginal
spines; t, tegmentum; vs, ventral scales) (after Maile, 1981).

electrode at the very first contact, general pressure on the
girdle, instead of the touch of single elements in the girdle,
can be excluded as a cause of the observed reactions, e.g.
avoidance behavior. The reactions were classified into four
categories of increasing intensity: movement of touched ele-
ment; movement of neighbouring elements; girdle movement
near the touched place; whole animal moving away. The fre-
quencies of these avoidance behavior patterns were deter-
mined by direct observation. Each of 20 animals was tested
several times (depending on the overall activity, that can make
the tests quite difficult) and in all areas (dorsal scale, marginal
spine, hair, ventral scale). The chitons were light-adapted for
at least one hour [dark-adapted animals exhibit a marked
response to light (Bergmann, 1986)]. The chitons had been

kept in aquaria containing artificial sea water (20°C, natural
day/night-cycle, simulation of high and low tides) for 6 months
up to 3 years before the tests.

RESULTS
MORPHOLOGY OF THE GIRDLE

The girdle of Chiton olivaceus is covered with
characteristic calcareous parts (Fig. 1). On the dorsal surface,
large scales are arranged like tiles on a roof, with the open
side directed towards the shell. The tallest scales (up to
180x130 «m) are found in the middle part of the girdle, whereas
those near the girdle margin and near the shell valves are
much smaller (50x20 um). In most cases, the surface of the
scales shows irregular elevations and ridges (Figs. 2, 3). The
girdle margin is formed by one row of calcareous spines
(50-90 um long and 20-25 um wide). They bear thin ridges
running roughly parallel to their long axes (Fig. 2). Between
the marginal spines and the dorsal scales, hair-like forma-
tions can be observed at regular distances. One hair is nor-
mally accompanied by one or two clapper-like structures. Each
hair or clapper consists of a solid shaft of organic material
and a calcareous tip, which can be lost in some hairs. The
hair shaft is 70-110 nm long and between 5 um (at the base)
and 1.5 um wide (distally). The calcareous tip structure can
reach a length of 30 um and a diameter up to 4 um. The
organic shaft of the clappers is 10 um long and 2 um wide;
the calcareous tip is 10-15 nm long and about 7 um in width.

Fig. 2. Scanning electron micrograph of the girdle margin (cl, clapper; cti, calcareous tip of a hair or clapper; c, cuticle; ds, dorsal scale;
h, hair; ms, marginal spine). Fig. 3. Isolated dorsal scale, KOH-treated. A groove (arrow) shows the connection site with the spicule cell of
the papilla. Fig. 4. Ventral scales (right side is lateral).

FISCHER ET AL.: GIRDLE SENSE ORGANS IN CHITON OLIVACEUS 133

In living animals, the hairs are straight and oriented at an angle
between 0° (parallel to the substratum) and 60°. Ventrally, the
girdle is covered with rows of small scales (30-40 xm long,
about 13 um broad) (Fig. 4). The rows are oriented perpen-
dicular to the girdle’s margin. All scales, at least at their base,
are embedded in the cuticle.

AVOIDANCE BEHAVIOR OF LIVING ANIMALS

We studied qualitatively the avoidance behavior of 20
individuals of different ages (age can be estimated from the
size of the animals). There was no difference in the reactions
between these chitons.

Generally, there is a reaction to touch of any calcareous
element but of a varying degree (Table 1). Girdle movements
in the stimulated region can be observed upon touch of every
structure. However, the weak reaction of the dorsal scales and
the hairs could be accidental, as girdle movements can
sometimes be registered without obvious external stimulation.
Individual ventral scales are not moved. They are embedded
in the cuticle except at their distal surface. The most effec-
tive stimulation is contact of a marginal spine. The touched
spine is moved away immediately with a subsequent move-
ment of neighboring spines. The girdle in the stimulated areas
is withdrawn and, after repeated stimulation the animal often
moves away. The reaction of the other structures to touch is
much weaker; the weakest is that of the hairs.

Because the clappers are very small and inserted very
close to the hairs, it was not possible to stimulate this struc-
ture without also possibly stimulating a hair. Therefore, the
clappers were omitted in Table 1.

PARENCHYME OF THE GIRDLE

The parenchyme of the girdle consists of a network of
connective tissue (Fig. 5). The nuclei of these cells are relative-
ly small. The space between the cells is filled with irregular
groups of collagen fibers and scattered muscle ceils. The mus-

Table 1. Avoidance behavior of Chiton olivaceus upon touch of single
elements in the girdle with a glass microelectrode (— = no reaction
observed, + = up to 30% of the tests were positive, ++ = 30-60%
of the tests were positive, +++ = > 60% of the tests were positive).
The ventral scales tested were in girdle areas which were not com-
pletely attached to the substratrum. Altogether, 60 tests were per-
formed for each of the girdle elements, except for the ventral scales
(31 tests).

movement movement

of of neigh- animal
touched boring girdle moves
element elements movement away
dorsal
scale + - + -
marginal
spine +++ ++ +4+4+ ++
hair - - + -
ventral
scale = - ++ =

cle cells insert at the basal lamina of the epidermis and not
at the hard structures of the girdle. The movement of spines
is obviously produced indirectly by contraction of the underly-
ing muscle cells. Hemolymph filled lacunae of various sizes
are bordered only partially by the cells of the connective
tissue; they continue to a large extent into the intercellular
substance.

EPIDERMIS

The girdle epidermis consists of 2.5-4.5 um high cells
that are intensively interdigitated. Distally, the epidermal cells
are connected by zonulae adhaerens and septate junctions.
Short microvilli (0.5-1 wm in length) protrude into the cuticle.
The nucleus fills most of the cell’s volume; its chromatin is
highly condensed, a sign of relatively low metabolic activity.
Many tonofilaments run from the basal lamina up to the tips
of the microvilli (Fig. 6). Scattered ribosomes and only a few
mitochondria are also found randomly in the epidermal cells.
In areas where new papillae are formed, epidermal cells show
ultrastructural features indicating higher metabolic activity.
They have a larger cell volume, the chromatin is less con-
densed and the cytoplasm contains more mitochondria and
some endoplasmic reticulum (ER). There is a regular transi-
tion to the secretory cell type of the papillae.

GENERAL PATTERN OF THE GIRDLE PAPILLAE

Papillae of various sizes (according to the position on
the girdle) insert in the epithelial layer. Generally, a papilla
contains a varying number of secretory cells, one ciliary cell
and one spicule cell. The external appearance of the forma-
tions on the girdle looks very different (Fig. 2). However, the
composition of the papillae (which are connected with these
formations) is essentially the same. Therefore, a detailed
description of the cell types found in a papilla is next
described.

SECRETORY CELLS

Secretory cells, as well as all other cells of the papilla,
are interconnected in the same way as the epidermal cells.
Active secretory cells have a relatively large nucleus without
much condensed chromatin which normally lies near the
basal lamina. Granular ER, free ribosomes and numerous
mitochondria are a regular feature of the cytoplasm. Golgi ap-
parati are rare, although the cell is filled to a large extent with
membrane-bound secretory granules. Other granules, of vary-
ing electron density, and a few multivesicular bodies are also
found. In older papillae, especially at the ventral side of the
girdle, the secretory cells’ activity decreases and the
chromatin becomes more condensed (new papillae are
formed mainly near the shell and the girdle margin, dorsally
and ventrally the zone between these areas contains older
papillae except in places where a scale had been lost.

CILIARY CELL

The ciliary cell also shows the ultrastructural features of high
metabolic activity. The large nucleus is surrounded by
granular ER; many mitochondria, a few granules and multi-

134 AMER. MALAC. BULL. 6(1) (1988)

Fig. 5. Cross section through the girdle, ventral part. Two papillae (p) are connected by cups with their scales (ct, connective tissue; c, cuticle;
e, epidermis; scu, spicule cup; vs, ventral scale (decalcified) (left is lateral, upper side is ventral). Fig. 6. Epidermis on the dorsal side of the
girdle (bl, basal lamina; co, collagen; mv, microvilli; nu, nucleus; tf, tonofilaments; arrows indicate branches of the cup of a dorsal scale running
down between the microvili of the epidermal cells). Fig. 7. Distal part of a ventral papilla (c, cuticle; sc, spicule cell; scu, spicule cup). A cilium
protruding from the ciliary cell can be seen (arrow). This cell invaginates into the distal part of the spicule cell (double arrow).

vesicular bodies are also present. Distally, the cell is elongated
and protrudes up to the cuticle. Relatively large (0.5 um in
diameter) microvilli protrude into the cuticle. One cilium (9+2
structure) runs to the base of the spine, scale, clapper or hair
(in these sections we refer to all these formations as
“‘spicules’’) (Fig. 7). This cilium originates from a striated
rootlet that consists of several parts (Fig. 8). Branches of the
ciliary cell invaginate into other cells, especially into the distal
area of the spicule cell (Figs. 9, 10). Two centrioles are pre-
sent in this invagination. In longer papillae, the distal part of
the ciliary cell contains many microtubules. This zone then
resembles a dendrite.

SPICULE CELL

The spicule cell connects the spicule with the papilla.
Again, the nucleus is large and does not contain much con-
densed chromatin. Especially in the distal half of the cell
agranular ER, numerous microtubules and many mitochon-

dria are present. This zone is also characterized by the in-
vagination of the ciliary cell mentioned above. The mem-
branes of both cells are parallel to each other, and the spicule
cell forms a dense network of microfilaments around this part
of the ciliary cell (Fig. 9). Distally, the spicule cell bears
numerous microvilli that are connected with the organic cup
or shaft of the spicule (Fig. 11). The calcareous element is
placed in or on top of this organic structure. It does not con-
tain any cellular elements. In all parts of the girdle, the cilium
of the ciliary cell at the base of the spicule is oriented towards
the girdle margin, i.e. the papillae are polarized. Structures
resembling small neurons (fibers containing numerous
microtubles) can be found from the basal lamina far up into
the papilla. However, no synapse or direct connection to a
cell could be seen so far.

VENTRAL PAPILLAE
The ventral papillae are oriented at an angle of 10-20°

FISCHER ET AL.: GIRDLE SENSE ORGANS IN CHITON OLIVACEUS 135

towards the girdle margin (Figs. 5, 12). A papilla consists of
about seven cells (one spicule cell, one ciliary cell and about
five secretory cells). The cup of the ventral scale consists of
three zones. The proximal filaments are very thin (about
15 nm; in the median zone these filaments become thicker
(150 nm). The area adjacent to the calcareous scale is
homogeneous and surrounds the scale continuously in
younger scales; in older ones the distal parts of the organic
component has been eroded. Newly formed scales lie near
the papillae, older ones have moved far into the cuticle. In
these papillae the ciliary and spicule cells are elongated up
to the scales’ cup. Finally, the scale is dropped and a new
one is formed (for a description of the formation of spicules
see Haas and Kriesten, 19795).

DORSAL PAPILLAE

The size of the papillae on the dorsal side of the gir-
dle varies considerably. They are small near the margin and

the shell valves, where the scales are also small, and very
large in the median area of the girdle. In this median area
one cannot clearly distinguish between distinct papillae; a
large scale can be surrounded by a ring of papillae-like cell
complexes. Only on one side, towards the shell valve, are the
spicule cell and the ciliary cell present (Fig. 11). Underneath
the scale, normal epidermal cells are present (Fig. 13). All
other cells are of the secretory type (Fig. 14).

The cup of a dorsal scale is composed of two parts
(Fig. 15). In most cases the basal plate has a straight border
towards the calcareous element; at the lower side, short
branches run down between the microvilli of the epidermal
cells (Fig. 6). At the lateral side, the scale is covered with an
organic sheet that reaches the basal plate; there is often no
direct connection between these two parts. The lateral part
has a straight border towards the cuticle and many processes
into the calcareous element. In new dorsal scales the basal
plate is formed some time after the lateral part.

Fig. 8. Longitudinal section through the base of a cilium of the ciliary cell (ci, cilium; mv, microvilli of the ciliary cell; r, striated rootlet of the
cilium). Fig. 9. Distal part of a ventral papilla (c, cuticle; cc, ciliary cell; df, dense network of small fibers around the invagination of the ciliary
cell; sc, spicule cell; scu, spicule cup; sec, secretory cell; arrow indicates basal body). Fig. 10. Tips of the ciliary celi and the spicule cell
(ce, ciliary cell; iv, invagination of the ciliary cell into the spicule cell; mv, microvilli; r, striated rootlet of a cilium; sc, spicule cell; scu, spicule
cup). Fig. 11. Section through the receptive part of a dorsal papilla (sc, spicule cell; scu, spicule cup; arrow indicates invagination of the ciliary

cell into the spicule cell; double arrow indicates ciliary cell).

136 AMER. MALAC. BULL. 6(1) (1988)

Fig. 12. Schematic drawing of a ventral papilla with its scale. Left side is lateral, upper side is ventral (c, cuticle; cc, ciliary cell; e, epidermal

cell; n, neurite; sc, spicule cell; scu, spicule cup; vs, ventral scale;).

Sum

Fig. 13. Schematic drawing of a dorsal papilla complex with its scale [c, cuticle; cc, ciliary cell; ds, dorsal scale; e, epidermal cells; n, neurite;
p, papilla; sc, spicule cell; scu, spicule cup); arrow indicates the two parts of the cup of the dorsal scale are attached to each other (right

side is lateral).

GIRDLE MARGIN

The greatest variety of structures is found in the girdle
margin. The papillae of the marginal spines, of the hairs and
the clappers are in many cases not distinct. All cells at the
margin (except spicule and ciliary cells) resemble secretory
cells; there are no typical epidermal cells.

At the margin, the papillae lie in three rows: ventrally,
the papillae of the marginal spines; medially, the papillae of
the clappers; dorsally, the papillae of the hairs (Fig. 16). The
papilla of a marginal spine has numerous secretory granules
concentrated in the upper side, whereas the papilla of a clap-
per has most of these granules in its lower side. The cup of
the marginal spines consists of the same zones as in the ven-
tral scales. In the clappers, the cup has been transformed
into an elongated shaft, which is solid except near the tip of
the spicule cell. Around the distal part of the papilla and the

base of the shaft, a cortex of darkly stained granules is
embedded in the cuticle. The middle zone of the papilla of
a hair is quite narrow. All cells, spicule cells and ciliary cells
as well as secretory cells, have a thin diameter in this zone.
Numerous microtubles contribute to the dendritic appearance
(Fig. 17). The distal part of the papilla is swollen (Fig. 18).
Secretory cells are highly vacuolized around the spicule and
ciliary cells. The hair shaft is solid except at the connection
with the spicule cell. The distal (swollen) part and the base
of the shaft, as in the clappers, are surrounded by a granulate
cortex; in the hairs, it can protrude out of the cuticle for a short
distance.

DISCUSSION

Despite the very different external appearance of the
girdle formations, all papillae in Chiton olivaceus are of similar

FISCHER ET AL.: GIRDLE SENSE ORGANS IN CHITON OLIVACEUS 137

construction. They are composed of a varying number of
secretory cells that surround one spicule cell and one ciliary
cell. The ciliary cell invaginates into the spicule cell which
is highly specialized in this zone. The spicule cell is connected
with the organic cup of a calcareous structure. Both these
components vary considerably in size. The same pattern is
also found in the primitive polyplacophoran Lepidopleurus ca-
jetanus Poli (Fischer, unpublished) as well as in Lepidochitona
cinereus (Haas and Kriesten, 1975). In Acanthochitona
fascicularis a similar appearance has been found (Fischer et
al., 1980) with three major differences: the secretory cells are
more prominent; photoreceptor cells are present in many
papillae; a stalked nodule protrudes from many papillae into
the cuticle. This nodule resembles the swelling of the hair
papilla in Chiton olivaceus. It looks like a distal part of a papilla
that has lost its spicule. In young Mopalia muscosa a pattern
similar to Chiton olivaceus is found (Leise, 1986) (Fig. 6). It
seems that the type described here is the basic structure of
the girdle sense organs in the polyplacophora.

In Acanthochitona fascicularis, another type of spine
has also been described, in addition to this general type.
These spines are not connected with a papilla. Each is based
on top of a large cup-like cell in the epidermal layer and grows
basally as the animal gets larger. In contrast, the ‘‘normal”’
type of spine does not grow after it is produced. Behavioral
observations and the fine structure of the cup-like cell sug-
gest that this spine type in A. fascicularis is merely defensive
(Fischer, 1979).

Adult mopaliid chitons have elaborate sensory hairs
in the girdle (Leise and Cloney, 1982; Leise, 1986). Leise (1986)
has demonstrated that these hairs are formed by the growth
of several spines (very similar to the hairs of Chiton) close
to one another. As they grow, the whole bundle is surround-
ed by an organic cortex. Thus, the complex hair in Mopalia
is an elaboration of the ‘‘normal”’ type.

In Acanthochitona fascicularis the spicule cell forms
a neurite (Fischer et a/., 1980). Nerves have also been
demonstrated in the girdle sense organs of Mopalia muscosa
(Leise and Cloney, 1982; Leise, 1986). In Chiton olivaceus,

structures resembling neurons are present in the papillae of
every type of girdle formation. However, the presence of such
structures seen in the electron microscope is only an indica-
tion of a sensory function, for two reasons. Cells that are
not sensory, such as the secretory cells in the aesthetes, can
form fiber-like extensions that are very similar in structure
to neurons. However, they certainly have another function, as
they are not connected with the nervous system (Knorre,
1925). If the fibers observed in the papillae are nerves, they
could have other functions such as stimulating the secretory
cells. To establish a sensory function, appropriate
neurophysiological or behavioral experiments must be car-
ried out. Neurophysiology in chitons is very difficult, as single
nerve fibers are thin and the amplitude of potential changes
is quite low (Fischer et a/., unpub. data).

The results of the stimulation experiments show that
the girdle sense organs are mechanoreceptors. Due to the
fact that the basic structure is the same in all species and
in all areas in Chiton, we suggest that, apart from the func-
tion of the secretory cells, mechanoreception is the basic func-
tion of the girdle papillae. A possible function of the secretory
cells could be to produce or impregnate the cuticle. The
chemical composition of the secretory granules is unknown.

The presence of mechanoreceptors is certainly of great
importance for a relatively small animal which lives in the tidal
region and moves actively, but slowly, on exposed substrata
for feeding. Individuals of Chiton olivaceus which have been
detached from their stone have great difficulty settling again
in turbulent water (pers. obs.). Under normal conditions, the
girdle is pressed onto the substratum. There is no gap and
the animals are not vulnerable to strong water movement. The
ventral scales could provide feedback information about the
pressure of the girdle on the substratum. The reactions to
stimulation of the marginal spines show that these structures
can detect an obstacle or movements of other animals.

Most chitons including Chiton olivaceus do not possess
eyes. The photoreceptor cells in the aesthetes (Fischer, 1978)
are involved in the photonegative behavior (Arey and Crozier,
1919; Boyle, 1972; Bergmann, 1984) and in the shadow

Fig. 14. Section through a dorsal papilla showing secretory cells (ct, connective tissue). Fig. 15. Cross section through a part of the cup of
a dorsal scale. The border between the basal plate (left) and the lateral part (right) is clearly seen (arrows) [ds, dorsal scale (decalcified)].

138

10 um

AMER. MALAC. BULL. 6(1) (1988)

Fig. 16. Schematic drawing of the margin of the girdle [c, cuticle; cl, clapper; clp, papilla of a clapper; cls, clapper shaft; co, cortex-like struc-
ture; cti,, calcareous tip of a hair or a clapper; e, epidermis; hs, hair shaft (a specialized spicule cup); h, hair; hp, papilla of a hair; ms, marginal

spine; msp, papilla of a marginal spine; scu, spicule cup].

response (Crozier and Arey, 1918). As mainly nocturnal
animals, the light sense is not very specialized in chitons. C.
olivaceus is frequently found on very irregular substratum.
They hide in small holes (such as produced by the clam
Lithophaga lithophaga L.) in stones. The great number of
lateral mechanoreceptors obviously is involved in orientation.

Stimulation of the hairs does not evoke a strong reaction,
touch apparently being not the appropriate stimulus. When the
animal is feeding, the hairs are oriented towards the open
water and are moved slightly by water motions (pers. obs.).
A possible function could be to measure these movements.

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$

The large dorsal scales certainly protect the animal
against predators or strong water movement. Most predators
usually only consume the foot and the viscera, not the valves
and the girdle (Leise, 1986). When disturbed, Chiton olivaceus
presses the girdle to the substratum very tightly. It is difficult
to detach the animals. For nearly all possible predators, this
species is unattractive because of the protection afforded by
the valves and the dorsal scales. However, the dorsal papillae
still retain the sensory elements, although they are relatively
small. Further experiments must be carried out to define the
exact function of the girdle sense organs of chitons.

«

oo

Fig. 17. Dendrite-like appearance of different cells (arrows) in the base of the distal part of hair papilla. Fig. 18. Distal part of a hair papilla
[cc, ciliary cell; sc, spicule cell; scu, spicule cup (here transformed into the shaft of the hair); vw, vacuolated secretory wall cells].

FISCHER ET AL.: GIRDLE SENSE ORGANS IN CHITON OLIVACEUS 139

ACKNOWLEDGMENTS

We thank Birgit Seibel for the final drawings. Roland Gantner
and Ulrike Kaltenhauser performed some preliminary studies. We are
grateful to Renate Kammerer for reading the manuscript. Two
anonymous reviewers provided useful comments. Last but not least,
we would like to thank Prof. G. A. Manley for the revision of our
English.

LITERATURE CITED

Arey, L. and W. J. Crozier. 1919. The sensory responses of Chiton.
Journal of Experimental Zoology 29:157-260.

Bergmann, W. 1986. das Verhalten von Chiton olivaceus
(Polyplacophora) bei gerichtetem Einfall von Licht.
Zulassungsarbeit. Technische Universitat Munchen. 80 pp.

Blumrich, T. 1891. Das Integument der Chitonen. Zeitschrift wissen-
schaftliche Zoologie 52:404-476.

Boyle, P. 1972. The aesthetes of chitons. II. Role in the light response
of the whole animal. Marine Behavioural Physiology 1:171-184.

Crozier, W. J. and L. Arey. 1918. On the significance of the reaction
to shading in chiton. American Journal of Physiology
46:487-492.

Fischer, F. P. 1978. Photoreceptor cells in chiton aesthetes (Mollusca,
Polyplacophora). Spixiana 1:209-213.

Fischer, F. P., W. Maile and M. Renner. 1980. Die Mantelpapillen und
Stacheln von Acanthochitona fascicularis L. (Mollusca: Poly-
placophora). Zoomorphologie 94:121-131.

Haas, W. and K. Kriesten. 1975. Studien uber das Perinotum-Epithel
und die Bildung der Kalkstacheln von Lepidochitona cinereus.
Biomineralisation 8:92-107.

Knorre, H. v. 1925. Die Schale und die Ruckensinnesorgane von
Trachydermon (Chiton) cinereus L. und die ceylonischen
Chitonen der Sammlung Plate. Jenaer Zeitschrift fur Medizin
und Naturwissenschaften 61:469-632.

Leise, E. 1986. Chiton integument: Development of sensory organs
in juvenile Mopalia muscosa. Journal of Morphology 189:71-87.

Leise, E. and R. Cloney. 1982. Chiton integument: Ultrastructure of
the sensory hairs of Mopalia muscosa (Mollusca: Polyplaco-
phora). Cell and Tissue Research 223:43-59.

Leloup, E. and P. Volz. 1938. Die Chitonen (Polyplacophoren) der
Adria. Thalassia 2, 10:1-63.

Maile, W. 1981. Drusenzellen und Drusenzellen-komplexe an Mantel,
Fuss und Kiemenrinne dreier Polyplacophoren-Arten.
Diplomarbeit, Universitat Munchen. 136 pp.

Moseley, H. N. 1884. On the presence of eyes and other sense organs
in the shells of the Chitonidae. Annual Magazine of Natural
History (series 5) 14:141-147.

Plate, L. H. 1898. Die Anatomie und Phylogenie der Chitonen, Teil
A. Zoologisches Jahrbuch, Supplement 4:1-241.

Plate, L. H. 1902. Die Anatomie und Phylogenie der Chitonen, Teil
B und C. Zoologisches Jahrbuch, Supplement 5:15-216,
281-600.

Reynolds, F. S. 1963. The use of lead citrate at high pH as an elec-
tron opaque stain in electron microscopy. Journal of Cell Biology
17:208-212.

Date of manuscript acceptance: 16 October 1987

SENSORY ORGANS IN THE HAIRY GIRDLES OF
SOME MOPALIID CHITONS

ESTHER M. LEISE'
DEPARTMENT OF ZOOLOGY
UNIVERSITY OF WASHINGTON
SEATTLE, WASHINGTON 98195, U. S. A.
and
FRIDAY HARBOR LABORATORIES
FRIDAY HARBOR, WASHINGTON 98250, U. S. A.

ABSTRACT

The polyplacophoran mantle secretes the shell plates, houses the gills in the pallial grooves,
and forms a muscular perinotum or girdle that encircles the shell and viscera. The epidermis of this
girdle occurs as papillae of columnar cells dispersed over an otherwise cuboidal epithelium. Depend-
ing upon the species, these papillae can produce a variety of hard structures: calcareous scales, spicules,
or spines and/or chitinous hairs. Some papillae also produce bulbous outgrowths called nodules or
‘“‘morgensternformigen Korper’’ (morning star-shaped bodies). These nodules contain the dendrites
of sensory neurons and are thought to be mechanoreceptive. Nodules can occur alone in the cuticle
or in conjunction with calcareous spicules. Nodules of this type are present in the hairs of chitons
in the genus Mopalia. Hairs from other mopaliid genera are also innervated, although they can lack
these particular structures. In most species of chitons that | examined, nodules are made in conjunc-
tion with the ventral girdle spicules and the marginal spicules. These presumptive mechanoreceptors
could be ubiquitous among chitons, as all species possess marginal spicules and overlapping ventral
spicules. Hairs could have evolved to extend the reach of these tactile receptors beyond the surface
of the animal’s body, as well as to provide mechanical protection from desiccation and predation.

The external surfaces of the polyplacophoran girdle are
armed with diverse types of secreted structures whose form
and arrangement is species specific. These secretions include
calcareous spicules, spines, and scales, and chitinous hairs
(Fischer-Piette and Franc, 1960) (Figs. 1, 2). The dorsal sur-
face can produce several types of hard parts, while the man-
tle edge and ventral surfaces generally produce one type of
ornament each (Hyman, 1967). These structures can be com-
pletely or partially embedded in the cuticle that covers the
epidermal cells of the girdle. These girdle formations, or orna-
ments, can be simple or composite structures (Fig. 1). In-
dividual, fusiform, calcareous spicules are often totally
embedded in the cuticle, which is 25 to 100 .m thick, whereas
longer calcareous spines (Figs. 1, 2b) have only their prox-
imal ends in the cuticular matrix (Plate, 1898, 1902; Hyman,
1967). Many species produce overlapping calcareous scales
(Fig. 2a) that are also connected to the cuticle basally. Species

1Present address: Otology Lab, 1159 Surge Ill, University of Cali-
fornia, Davis, CA 95616.

in several families produce hairs (Fig. 2c), often called setae
or bristles, that can be simple, jointed (articulated), or com-
posite chitinous shafts that extend beyond the girdle surface.
Hairs usually consist of an extension of the cuticular matrix
and can be surrounded by a more densely staining cortex
(Leise and Cloney, 1982).

Most spicules are surrounded by a layer or ‘‘cup’”’ of
material that is darker than the enveloping cuticular matrix
and stains more densely in sectioned material (Figs. 1, 3)
(Plate, 1898, 1902; Knorre, 1925; Leise and Cloney, 1982). In
spicules from many species, this dense cup is elongated in-
to a shaft that extends from the spicule to the epidermal cells
(Fig. 1). The similarity of many hairs to this type of spicule
shaft and the presence of a spicule at the distal tip of many
hairs, led Thiele (1929) and Hyman (1967) to suggest that
spicules and hairs represent the two ends of a continuum of
girdle structures. They regard hairs as highly modified shafts
of spicules. | continue their usage here and refer to hairs as
those structures in which a chitinous shaft projects above the
surface of the girdle and is the predominant part of the organ.

American Malacological Bulletin, Vol. 6(1) (1988):141-151

144

142 AMER. MALAC. BULL. 6(1) (1988)

50 um

SPINE

DISTAL
NODULE SHAFT

CUTICLE ANNULUS

SNe —~ Gases <
PIGMENT GRANULES PROXIMAL SHAFT

Fig. 1. Diagram of five types of spicules and one spine. a. Primary
spicule from a newly metamorphosed juvenile. Note thin chitinous
cup. b. Spicule with apically and basally thick cup. c. Spicule with
pigment granules and shaft. d. Spicule with an annulate shaft sur-
mounting a sensory nodule. e. Spicule as in d but with an articulated
shaft (from Leise, 1983).

As will be described below, most hairs contain or are
in contact with dendrites from presumptive sensory neurons.
This paper reviews the morphology of chiton hairs while focus-
ing on their neuronal elements and describes the relation-
ships of these hairs to other girdle ornaments.

DIVERSE GIRDLE HAIRS: AN OVERVIEW

Hairs occur in a bewildering range of sizes and con-
figurations in species from at least five families: Chitonidae;
Lepidochitonidae (Ferreira, 1982); Callochitonidae; Chaeto-

pleuridae; and Mopaliidae [classification after Bergenhayn
(1955) unless otherwise cited]. In addition, hairs from many
species of chitons will erode during the animal’s lifetime. Thus,
it can be difficult to understand the morphology of a particular
type of hair if only large hairs or hairs from old animals are
studied. Species such as Chiton olivaceus Spengler, 1797 (fam-
ily Chitonidae) can produce small marginal hairs 80 to 100 nm
long (Plate, 1902). In the Lepidochitonidae (Ferreira, 1982),
species such as Tonicella insignis Reeve, 1847 produce small,
simple hairs only 100 xm long (Leise, 1983), while others, such
as Dendrochiton lirulatus Berry, 1963, produce tufts of hairs
up to 500 um long. Hairs from species of Callochitonidae, such
as Eudoxochiton nobilis Gray, 1843, often have large ar-
ticulated shafts about 1.5 mm in length (Leise, 1983). On in-
tact animals of E. nobilis, even the distal spicules can be
discerned. Species in the Chaetopleuridae and Mopaliidae
also display hairs in a wide range of sizes; although the
Chaetopleuridae characteristically produce hairs (Pilsbry,
1893), some species, like Chaetopleura lurida (Sowerby, 1832)
secrete none. The girdle of this species bears spicules with
articulated and simple shafts. A congener, C. peruviani
Lamarck, produces similar spicules whose elongated shafts
extend beyond the cuticular surface and so earn them the
designation of hair (Plate, 1902; Fischer-Piette and Franc,
1960) (Fig. 4). Among the Mopaliidae are also species that
produce small, simple hairs, such as those on Katharina
tunicata Wood, 1815 or very large, simple hairs, as are found
on Plaxiphora obtecta (Carpenter in Pilsbry, 1893) (Table 1).

Most of the above mentioned hairs conform to the
hypothesis of Thiele (1929) and Hyman (1967) that hairs are
elongated spicule shafts. However, the large hairs secreted
by species in the genera Mopalia and Placiphorella, and those
secreted by some of the Lepidochitonidae, namely Lepido-
chitona flectens (Carpenter, 1864), and species in the genus
Dendarochiton Berry, 1911 (Ferreira, 1982), do not conform to
Thiele’s (1929) and Hyman’s (1967) hypothesis. These latter
types of hairs are composite structures, built by the replica-
tion of many basic units. They are not simply enlarged or
elongated spicule shafts. In the genus Mopalia, the basic unit
construct is a calcareous spicule and its long chitinous shaft.
This basic unit is serially repeated along an outgrowth of the
cuticle, and with the exception of the groove along which these
spicules lie, the entire organ is surrounded by one or two
distinct layers of dense cortical material (Fig. 5) (Leloup, 1942;

Table 1. Characteristics of chitons hairs in the family Mopaliidae (Structure: C = compound; S = sim-
ple and lacking medulla. Length and width are maxima recorded. Cortex: ++ = >20 um thick; +

= <20 um thick; - = lacking).
Species Hair Length Hair Width Structure Cortex Innervation
(mm) (um)

Mopalia muscosa 5 400 C ++ +

M. ciliata 3 200 Cc ++ +

M. lignosa 3 300 Cc + +

M. hindsii 25 80 C + +
Plaxiphora obtecta 2 300 S ++ -
Katharina tunicata 0.1 5 S) + +
Placiphorella velata 5 400 Cc - +

LEISE: CHITON SENSORY HAIRS 143

Leise and Cloney, 1982). Similarly, in the genus Placiphorella,
the hair is an extension of the cuticle and is entirely covered
with spicules that lie in whorls just below the surface of the
hair (Fig. 2c) (Plate, 1902). From Ferreira’s (1982) descriptions,
the hairs of L. flectens and the genus Dendrochiton appear
likewise to be branched or compound structures and not simp-
ly enlarged spicule shafts.

THE MORPHOLOGY OF HAIRS OF MOPALIA
MUSCOSA: A MODEL FOR COMPOSITE
SENSORY HAIRS

A fully-formed hair of Mopalia muscosa is a curved,
distally tapered extension of the cuticle that bears a mesial
groove in which lies a row of spicules (Figs. 5, 6). Each
spicule occurs atop a distinct shaft, whose proximal end is
embedded in the cuticular matrix, or medulla. The medulla
is enveloped by a bilayered cortex, except for the mesial
groove, and is therefore exposed to the environment along
the length of that groove. Within the medulla, the proximal
end of each spicule shaft surmounts a bulbous epidermal pro-
jection, a stalked nodule (Leise and Cloney, 1982) or
‘“‘morgensternformig Korper’’ (morning star-shaped body)
(Reincke, 1868). Blumrich (1891), Knorre (1925), and Plate
(1898, 1902) described such nodules in many species. All of
these authors suggest that the nodules are tactile. Until recent-
ly (Leise and Cloney, 1982), their presence in hairs of the

p , Ye

Fig. 2. a. Dorsal integument of Lepidozona cooperi (Pilsbry, 1892) demonstrating overlapping scales. b. Dorsal integument of Acanthopleura
granulata (Gmelin, 1791) displaying calcareous spines. Cuticle is visible between spines. c. Dorsal hairs of Placiphorella velata. Numerous
spicules (arrows) are embedded near the surface of each hair (cu, cuticle; sh, shell) (from Leise, 1983). Fig. 3. Transverse 1 wm section through
the decalcified integument of Mopalia muscosa. Spicules (s) produced by spiniferous papillae (sp) contain brown pigment granules (arrow).
One spiniferous papilla has produced a sensory nodule (no). Part of its stalk is not in the plane of this section. Common epidermal cells (cec)
occur between papillae (from Leise and Cloney, 1982).

Mopaliidae was unknown.

The dorsal girdle epidermis is a single layer of cells
that is divided into numerous packets or papillae of colum-
nar cells. These papillae produce the hairs, spicules, and
nodules. Smaller cuboidal cells occur ubiquitously between
the papillae. The papillae that produce the hairs are the largest
in the epidermis and as a hair matures, the papilla comes
to lie in a small depression or pocket below the level of the
rest of the epidermal cells (Leise and Cloney, 1982; Leise,
1986).

Each subcortical cell produces a bundle of cortical
fibers (Figs. 6, 7). The fiber bundles of the inner cortex are
more dense than those of the outer cortex (Leise and Cloney,
1982). Each layer of the cortex in a mature hair is several
bundles thick, whereas in young hairs the cortex is only one
bundle wide. Newly forming hairs have no cortex and start
as a single spicule with an elongated shaft that lies above
a stalked nodule. More spicules and their associated shafts
and nodules are added to the growing cuticular hair and on-
ly after several nodules are present does cortex begin to ap-
pear. The cortex is initially a narrow crescent along the lateral
edge of the hair. As development proceeds, the hair grows
longer and the cortex become progressively wider until it en-
compasses nearly the entire shaft (Leise, 1986).

Submediullary cells occur as a hillock that protrudes
into the base of the hair shaft and presumably secrete the
medullary matrix. The sensory cells lie in clusters within this

144 AMER. MALAC. BULL. 6(1) (1988)

CORTEX

MEDULLA

MESIAL SPICULE

DENDRITIC sai a

Mopa/lia muscosa 0.25 mm

Fig. 4. a-d. Diagram of spicules and hairs of Chaetop/eura peruviana. a. Three hairs (h) occur above the cuticle (cu). All three hairs are simple;
the right two are each capped by a spicule (s) and appear to contain stalked nodules (no). b. Tip of a simple hair. c. Tip of an articulated
hair with distal spicule (s). d. Enlargement of the small spicule circled in a whose shaft contains a stalked nodule (no) (de, dermis) (from Plate,
1902). Fig. 5. Diagram of the external morphology of a hair of Mopalia muscosa. The base of the shaft of each mesial spicule is embedded
in the medulla. Below each shaft is an epidermal sensory nodule. The cut ends of the nodule stalks are visible in the medulla. The hair is
drawn in its entirety as if it were cut off just beyond the cuticle (from Leise, 1986). Fig. 6. Diagrammatic longitudinal section through the base
of a hair of Mopalia muscosa, drawn passing through the mesial groove and two spicules (s). Dendrites from three sensory neurons terminate
in nodules (no). In mature hairs, the sensory neurons occur in clusters, not as single cells, as they are drawn here, for clarity. Two nerves
(ne) cross the basal lamina (bl) as they emerge from the base of the papilla (cec, common epidermal cells; cu, cuticle; g, pigmented glial
cells; ic, inner cortex; m, muscle fiber; me, medulla; oc, outer cortex).

LEISE: CHITON SENSORY HAIRS 145

hillock and each cluster produces a long bundle of dendrites
that extends through the hair (Figs. 6-9) (Leise and Cloney,
1982). The oldest dendritic bundle extends to the tip of the
hair; younger bundles are progressively shorter. Each den-
dritic bundle ends in a nodule, just below the shaft of a mesial
spicule (Fig. 6). A hair can have from one to 20 nodules in
it, arising from the same number of neuronal clusters in the
submediullary hillock (Leise and Cloney, 1982). One or several
nerves emerge from the base of each trichogenous (hair-
producing) papillae (Figs. 6, 7, 10). These nerves are pre-
sumed to contain the axons of the submedullary sensory
neurons (Fig. 10). Although these basal axons have not been
definitively shown to arise from the neurons (i.e. the
submedullary neurons could be axonless, synapsing upon
sensory interneurons from the CNS, or the submedullary
“‘neurons’”’ could have been misidentified and the nerves
could have other functions) (see also following section), the
most obvious explanation is that the epidermal cells whose
long apical necks contain numerous parallel microtubules are
primary sensory neurons (Leise and Cloney, 1982). Finally,
there are usually fewer nerves than nodules within one papilla,
indicating that the axons from several clusters of neurons con-
verge onto a single nerve (Fig. 6).

Each nodule (and hence each dendritic bundle) con-
tains dendrites from several cells, there being from one to 25
dendrites per bundle (Fig. 9) (Leise and Cloney, 1982). Each
bundle is surrounded by one or two submedullary support-
ing cells. The dendrites often branch, so a tally of the number
of dendrites in a bundle overestimates the number of sen-
sory neurons. In figure 6 the sensory dendrites are drawn as
straight cylinders with only one neuron per cluster for ease
of presentation. Within the nodule, the dendrites ramify be-
tween the processes of the submedullary supporting cells that
contain large vacuoles (Fig. 11).

SENSORY HAIRS FROM OTHER
MOPALIIDAE

To gain some understanding of the occurrence of sen-
sory hairs throughout the Mopaliidae, | examined the girdle
integuments of six other species in this family. Animals were
collected from rocky intertidal regions in Puget Sound,
Washington, or on Vancouver Island, British Columbia (Leise,
1983). Samples of girdle integuments were fixed in Millonig’s
phosphate buffered glutaraldehyde and post-fixed in bicar-
bonate buffered osmium tetroxide (Cloney and Florey, 1968).
Detailed procedures are described elsewhere (see Leise and
Cloney, 1982; Leise, 1983). Specimens of Plaxiphora obtecta
were obtained indirectly from New Zealand, where they were
fixed in 5% formalin in seawater.

In addition to various shell and body characteristics,
one of the mopaliid diagnostic features is the production of
dorsal girdle hairs. From most accounts, the one exception
in this hairy family appeared to be Katharina tunicata. However,
Leloup (1940) noticed that the girdle of this species produces
tiny translucent hairs (Table 1). | confirmed this observation
and found that the papillae that secrete these hairs are also
innervated (Fig. 12).

Three other species of Mopalia, namely M. ciliata, M.
hindsii, and M. lignosa, have innervated hairs similar to those
of M. muscosa (Fig. 13). Interspecific variation occurs in size,
number of nodules per hair, extent of cortical envelopment,
and size and arrangement of spicule shafts (Leise, 1983).

The hairs of Placiphorella velata Dall 1878 (Fig. 2c) are
quite different from those in the genus Mopalia. Placiphorella
hairs contain no nodules, although they are innervated (Plate,
1902; Leise, 1983). Instead of lying above a nodule, each
spicule in these hairs lies above a cell that projects beyond
the hillock on a thin stalk (Fig. 14). The ultrastructure of these
cells deserves attention as they too are likely to be sensory
neurons. As Plate (1902) reported for P stimpsoni (Gould,
1859), several nerves emerge from the epidermis below each
of the hairs of P velata. Again, these nerves probably carry
axons from the primary sensory neurons, and axons from
many neurons converge into each nerve.

| also examined the hairs of Plaxiphora obtecta, which
are large discrete shafts of cortical material (Fig. 15, Table 1).
In sectioned material | found no nerves emerging from the
bases of their trichogenous papillae. With this exception, all
of the mopaliid hairs that | examined either contained or con-
tacted epidermal neurons (Leise and Cloney, 1982; Leise,
1983). The hairs of P obtecta could truly lack innervation, or
this lack could be the result of inadequate fixation.

Stalked nodules, such as those in hairs of mopaliid
genera, have been observed in the epidermis of many chitons
(Fig. 3; Table 2) and repeatedly hypothesized to be tactile
(Blumrich, 1891; Plate, 1898, 1902; Knorre, 1925; Thiele, 1929;
Haas and Kriesten, 1975; Fischer et a/., 1980). However, the
papillae that produce these nodules had not been shown to
send nerves into the dermis until the work of Leise and Cloney
(1982; Leise, 1983). All stalked nodules are not identical, as
is discussed below. The functional distinctions between the
various types of nodules are unknown.

OCCURRENCE OF SENSORY NODULES
IN THE CHITONS

According to Blumrich (1891), all chitons possess a
fringe of spicules around the mantle edge. In many cases,
the shafts of these marginal spicules contain or surmount a
stalked nodule (Table 2) (Plate, 1898, 1902; Knorre, 1925). The
hollow shafts of spicules in some species contain more
claviform (club-shaped) cellular protrusions that lack a slender
stalk (Blumrich, 1891; Plate, 1898, 1902; Knorre, 1925). | ex-
amined the ultrastructure of claviform nodules in Katharina
tunicata and found that they too contain dendrites from epider-
mal sensory neurons and that the dendrites ramify between
vacuolated processes of epidermal supporting cells. Other
epidermal protruberances described by Fischer et a/. (1980)
resemble incipient stalked nodules of Mopalia muscosa (Leise,
1983). In this review | refer to all of these epidermal protru-
sions as stalked nodules.

Only on the dorsal surface of the girdle are stalked
nodules reported to occur alone (Fig. 3) (Blumrich, 1891; Haas
and Kriesten, 1975; Fischer et a/., 1980; Leise and Cloney,

146 AMER. MALAC. BULL. 6(1) (1988)

Fig. 7. Median longitudinal 1 wm section through the base of a girdle hair of Mopalia muscosa. Common epidermal cells (cec) line the pocket
in which the trichogenous papilla (tp) lies. This section grazes the shaft (asterisk) of a mesial spicule (c.f. Fig. 6). The continuity of the medulla
and cuticle is visible just above this shaft. The proximal end of the dendritic stalk of a sensory nodule is emerging from the papilla (arrowhead).
One nerve (ne) emerges from the base of the papilla and enters the dermis (de) (ic, inner cortex; oc, outer cortex) (from Leise and Cloney,
1982). Figs. 8. Transverse 1 um section through a hair of Mopalia muscosa, above the cuticle. Six dendritic bundles (arrowheads) and one
nodule (double arrowheads) lie in the medulla. The groove (gr) in the cortex exposes the medullary matrix to the environment and is broader
in younger hairs. The shaft (arrow) of the last mesial spicule lies just inside the groove (from Leise and Cloney, 1982). Fig. 9. Transverse
section through the stalk of a sensory nodule from a hair of Mopalia muscosa. Numerous dendrites (d) are enclosed by two supporting cell
(sc). The dendrites contain numerous parallel microtubules (arrows) and mitochondria (me, medulla). Fig. 10. Axons (a) emerge from the base
of a trichogenous papilla (tp). The nerve passes into the dermis (de) from the base of the papilla. Note that the epidermal basal lamina (bl)
does not surround the nerve.

LEISE: CHITON SENSORY HAIRS 147

Fig. 11. Electron micrograph of a sensory nodule from a hair of Mopalia muscosa. In nodules, the dendrites (d) lose their typical organization;
their mitochondria are twisted and the microtubules are no longer in parallel arrays (arrowheads). Large electron-lucent vacuoles (ve) lie around

the periphery within the surrounding cells.

1982; Leise, 1986). In most cases, dorsal nodules are subja-
cent to spicules. The ventral girdle in all chitons produces
overlapping spicules (Blumrich, 1891; Pilsbry, 1892, 1893;
Knorre, 1925; Fischer-Piette and Franc, 1960; Hyman, 1967)
and in many cases these spicules also contact sensory
nodules (Table 2). Two exceptions are Placiphorella velata and
P. stimpsoni, in which the ventral spicules contact stalked cells
that are much like those in the dorsal hairs. These cells too
will probably prove to be sensory neurons upon further study.
Curiously, in P velata the marginal spicules are associated
with typical stalked nodules (Plate, 1902; Leise, 1983).

Of the chitons | studied, in only two species did | find
claviform nodules without innervated papillae: Eudoxochiton
nobilis and Plaxiphora obtecta. These animals were fixed in
5% formalin (see Leise, 1983) which does not preserve cellular
ultrastructure as well as the combination of glutaraldehyde
and osmium tetroxide. Thus, it is possible that the slender
(1-2 um in diameter) epidermal nerves were not preserved well
enough for me to recognize them. It would be most surpris-
ing if these two species alone show no innervated epidermal
sensory organs.

FUNCTIONS OF CHITON HAIRS

The functions of chiton hairs are not well understood
although plausible hypotheses abound. Hyman (1967)
describes chiton hairs as armature, although chitons bear-
ing hairs are successfully preyed upon by starfish (Mauzey
et al., 1968; Paine, 1980), seagulls (Moore, 1975), fish (Ronald
Shimek, pers. comm.) and humans. The girdle could be tox-
ic or distasteful but it does not provide sufficient protection

Fig. 12. Longitudinal 1 um section through the base of a trichogenous
papilla (tp) of Katharina tunicata. One nerve (ne) emerges from the
base of the papilla then continues into the dermis (de) (from Leise,
1983). Many cells of these papillae also produce granules, which can
be seen here in their various stages of condensation. Eventually,
granules are extruded into the cuticle.

148 AMER. MALAC. BULL. 6(1) (1988)

Table 2. Location of sensory nodules in or in conjunction with the designated girdle ornament. Alone
indicates in cuticle without attached spicules or hairs [* = animals | examined. Superscripts 1, 2, and
3 designate information from Plate (1898, 1902); Knorre (1925) and Fischer et al., (1980), respectively.

NA = not applicable].

Family and Species Alone

Lepidopleuridae
Lepidopleurus cajetanus'
Ischnochitonidae
Ischnochiton herdmani2 -
Lepidozona retiporosus* -
Lepidochitonidae
Lepidochitona dentiens* +
L. cinerea? +
Tonicella insignis*
Callochitondae
Eudoxochiton nobilis* -
Chaetopleuridae
Chaetopleura peruviana' -
C. lurida* -
Mopaliidae
Plaxiphora obtecta* -
Katharina tunicata-
Katharina tunicata*
Mopalia ciliata*
M. lignosa*
M. muscosa*
Placiphorella velata* -
Chitonidae
Chiton olivaceus' -
Acanthochitonidae
Acanthochiton fascicularis3 -

+++ 4+ 1

against predation. Predators tend to eat the foot and viscera,
discarding the shell and girdle.

Species with large and abundant hairs such as Mopalia
muscosa often support extensive epiphytic and epifaunal com-
munities (Phillips, 1972). This covering retains water and could
protect the animal against desiccation at low tides. This cover-
ing could also provide an additional defense against preda-
tion. Pisaster ochraceus (Brandt, 1835) will feed on M.
muscosa, but if the chiton is covered with its normal detrital
cloak, the starfish may fail to recognize it. After it touches
an overgrown chiton, a starfish will ignore it. The basis for
this protection, that is, whether the starfish’s olfactory or tac-
tile senses are deceived, is unknown. If the starfish contacts
the girdle of a clean chiton, it detects a prey item and removes
the chiton from the substratum. A chiton cannot escape a
hungry starfish nor maintain a sufficiently strong grip on the
substratum to avoid being consumed (pers. obs.). A chiton’s
epiphytic cloak could also afford protection from visual
predators. Chitons with well developed epiphytic communities
often resemble clumps of algae. Even during high tides, while
they are moving and feeding, their identity could be con-
cealed, as their slow rate of motion does not reveal their
animal nature.

In addition to providing passive defenses, chiton hairs
also mediate active responses from the animal. Chitons whose
hairs are bent or pinched will turn away from the source of

Dorsal Marginal Ventral Dorsal
Spicules Spicules Spicules Spicules
+
+ + NA
- + + NA
- + NA
NA
- +
- +
+ + + NA
NA + -
- + + +
- +
- + + +
- + + +
-_ + _ _
NA + +
+ - + NA

stimulation, or after several stimuli, tighten their grip on the
substratum and remain motionless. This response appears
to habituate rapidly, as prolonged or repeated stimulation will
soon fail to invoke a response (Leise, 1983).

This tactile aspect of hair function could be most im-
portant to juveniles. In Mopalis muscosa, hairs first appear
at metamorphosis (Leise, 1984) and although they do not in-
itially display all of the adult characteristics, the first sensory
neurons have differentiated and are presumably operational
(Leise, 1986). These young animals take refuge in cracks and
crevices in the substratum and their hairs may be impor-
tant detectors of irregular surface features. Similarly, ventral
nodules, which are widespread among the chitons, would give
an animal feedback on the surface characteristics of its
substratum and allow it to modulate its grip.

Although chiton hairs respond to touch, mechanore-
ception may not be their primary function. For example, they
could be chemoreceptive. However, unlike other molluscan
chemoreceptors (Laverack, 1968), the dendrites in the stalked
nodules are embedded in the cuticle. | found no pores in the
cuticle as exist in insect chemoreceptive hairs (Laverack,
1968). | was also unable to elicit any response from Mopalia
muscosa upon application to the hairs (without moving the
hairs) of various algae or tube feet from a predator starfish,
Pisaster ochraceus.

As previously stated, a sensory function is the most

LEISE: CHITON SENSORY HAIRS 149

MESIAL SPICULES

CORTEX CORTEX

CORTEX

MEDULLA

> MESIAL SPICULES

MEDULLA

DENDRITIC faa MEDULLA
, ; DENDRITIC BUNDLE, ANIAY
a
/ DENDRITIC BUNDLE
Mopalia ciliata 0.25 mm 0.50 mm Mopalia hindsit Mopalia lignosa 0.25 mm

Fig. 13. Diagrams of the external morphology of hairs from three species of Mopalia. Note that the cortex does not enclose the medulla in
hairs of M. hindsii (from Leise, 1983).

Fig. 14. a. Diagram of a hair of Placiphorella stimpsoni (from Plate,

1902). Note spicules (s) atop individual cells (arrows) beyond the

hillock of submedullary cells. One nerve (ne) emerges basally.

Inset: micrograph of a similar spicule and its subjacent cell from

a hair of P velata. b. Longitudinal 1 um section through the base

of a partially decalcified trichogenous papilla of P velata show-

ing a nerve (ne) emerging at the base (de, dermis) (from Leise,

1983). Fig. 15. Longitudinal 1 zm section through two dorsal hairs

(h) of Plaxiphora obtecta in one epidermal invagination. The right

hair shaft is still being formed, while the extrusion of the left hair

-—*. has ceased. Its papilla has produced a claviform cellular pro-
15 trusion below the shaft. No nerves have been found to emerge

: from these papillae (cu, cuticle; de, dermis) (from Leise, 1983).

150 AMER. MALAC. BULL. 6(1) (1988)

parsimonious explanation for the presence of an innvervated
integument and cells that resemble sensory neurons.
However, this explanation does not exclude the possibility that
the basal nerves mediate other functions, such as contrac-
tion or secretion. | found no obvious contractile elements in
the epidermis of Mopalis muscosa, although its skin does
secrete the cuticle and ornaments. Epidermal cells in other
species such as Katharina tunicata extrude pigment granules
into the cuticle (Fig. 12) (Leise, 1983). Whether or not the
nerves carry axons from neurons mediating epidermal secre-
tion is unknown.

CONCLUSIONS

My results lead me to suggest that most chiton hairs
are mechanoreceptors, although hairs are not the only girdle
sensory organ. Stalked nodules occur far more widely than
hairs, on the marginal and ventral surfaces of what may be
a majority of the chitons (Table 2). These nodules are pro-
bably important sources of feedback to the animal about the
nature of the surface on which it lives. Fischer et a/. (1980)
have also recognized photoreceptors in the girdle of Acantho-
chiton fascicularis that could in part be responsible for this
chiton’s response to changes in light intensity. Unfortunate-
ly, the existence of these girdle sensory organs is not widely
recognized.

In her review of the functional morphology of the chiton
epidermis, Hyman (1967) did not assimilate Plate’s (1902) in-
formation about the sensory nature of girdle hairs nor the sen-
timent from the German literature that stalked nodules are
tactile (Blumrich, 1891; Plate, 1898; Knorre, 1925; Thiele,
1929). Since then, the sensory nature of girdle structures has
been studied or remarked upon by several authors (Beedham
and Trueman, 1967; Haas and Kriesten, 1975; Fischer et al.,
1980). Most invertebrate texts include descriptions of chiton
sensory organs in the mouth, on the subradular organ, in the
buccal cavity, in the pallial grooves, and in the shell plates,
but not in the girdle (Hyman, 1967; Meglitch, 1971; Gardiner,
1972; Barnes, 1987; Pearse et a/., 1987).

In Mopalia muscosa, hairs erode and lose spicules
throughout the animals’s life. As many species produce hairs
and do so constantly during their lifetimes, the benefits from
their presence must outweigh their productive costs. Hairs
appear to have evolved several times in this class, as large
hairs occur in diverse families and can be formed in several
ways. Evolutionarily, there appear to be trends towards an in-
crease in the size of girdle ornaments (Pilsbry, 1892; Leise,
1983) and towards an inclusion of sensory organs in these
ornaments. Hairs are thus considered to be phylogenetically
advanced features, as they also occur in stratigraphically
newer families (Smith, 1960) and appear late in an animal’s
development.

The integument of most molluscs is richly endowed
with sensory organs and individual sensory neurons that serve
many modalities, including mechanoreception, chemorecep-
tion, and photoreception (Laverack, 1968). For the chitons to
be ‘“‘blind’’ to environmental stimuli over a large portion of
their skin would indeed be surprising (Beedham and Trueman,

1967). The work of many authors reviewed here suggests that
this is certainly not the case and that the girdle ornaments
are not just passive armature but active participants in the
lives of these animals.

ACKNOWLEDGMENTS

| thank Dr. Bradley R. Jones for his critical review of this
manuscript. This paper arose from a talk presented at the Symposium
on the Biology of the Polyplacophora at the 1987 meeting of the
American Malacological Union. | am also indebted to organizations
that supported this research: The Lerner-Gray Fund for Marine
Research of the Amerian Museum of Natural History, the Western
Society of Malacologists, Sigma Xi, and the Pacific Northwest Shell
Club.

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LEISE: CHITON SENSORY HAIRS 151

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Philadelphia, Pennsylvania.

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Co., Amsterdam.

Date of manuscript acceptance: 13 November 1987

THE ULTRASTRUCTURE OF THE AESTHETES IN LEPIDOPLEURUS
CAJETANUS (POLYPLACOPHORA: LEPIDOPLEURINA)

FRANZ PETER FISCHER
INSTITUT FUR ZOOLOGIE, TECHNISCHE UNIVERSITAT MUNCHEN
LICHTENBERGSTRASSE 4, D 8046 GARCHING,
FEDERAL REPUBLIC OF GERMANY

ABSTRACT

The aesthetes of Lepidopleurus cajetanus Poli consist of five different cell types: one or two
photoreceptor cells are present in the periphery in many of these organs. Products of tall secretory
cells pass through a perforated apical cap to the outside. Central cells probably are chemoreceptors.
Micraesthete cells form lateral branches from the main stem and end with unperforated caps at the
shell surface; their function is unknown. Peripheral cells form most of the border to the calcareous
shell substance. It is proposed that this is the general composition of the aesthetes in chitons.

Aesthetes are numerous organs in the upper shell layer
of the Polyplacophora (Figs. 1, 2). In recent years their
fine structure has been studied in several species. Except for
the species Acanthochitona fascicularis L. (Acanthochitonina)
(Fischer, 1979), only members of the Chitonina have so far
been examined in this respect (Boyle, 1974; Haas and
Kriesten, 1978; Fischer and Renner, 1978; Baxter et al., 1987).
For the discussion on the function of these unique organs
it is important to know which features are constant in the
aesthetes and which are species-specific variations. In the
present paper the aesthetes of a member of the relatively
primitive suborder Lepidopleurina are described and their
possible functions are discussed.

MATERIAL AND METHODS

Adult polyplacophorans of the species Lepidopleurus
cajetanus Poli were collected in the subtidal (about 1 m below
low tide level) region on the coast of northern Yugoslavia. Parts
of the tegmental shell layer containing the aesthetes were re-
moved and fixed in 5% glutaraldehyde in phosphate buffer
(pH 7.4) for two hours and postfixed in 2% osmium tetroxide
for two hours, all at 3°C. After dehydration in ethanol and pro-
pylene oxide the specimens were embedded in Durcupan.
Some of the specimens were decalcified overnight in chilled
3% EDTA in phosphate buffer after glutaraldehyde fixa-
tion. The others were split into two pieces and the calcareous
parts were removed by use of 5% HCl after embedding. Since
the tissue is already penetrated by the embedding material,
no damage occurred to the cells during this procedure, follow-
ing which the specimens again were embedded in Durcupan

to fill the holes left by the calcareous parts. Ultrathin sections
were cut with a LKB or a Reichert ultramicrotome, stained
with uranyl acetate and lead citrate (Reynolds, 1963) and
studied in a Zeiss or a Jeol electron microscope.

For scanning electron microscopy, the organic material
in the shell valves was removed by the use of concentrated
KOH at room temperature for about one hour, cleaned in an
ultrasonic cleaner and air dried. Other shells were air dried
without previous treatment. The specimens were given a 300
A thick coating with gold and were examined in a Cambridge
SEM.

RESULTS
SHELL SURFACE

The head valve has the shape of a half circle with a
few concentric ribs on the surface. In contrast, the other seven
valves show two different surface areas (Fig. 3): the lateral
fields resemble the head valve; parallel ribs are oriented along
the long axis of the animal in the second to the seventh valve
and are semicircular in the last one. In the median area of
the valves II-VIII, 60 wm-wide elevations form rows that run
mainly in the long axis. Parts of the articulamentum, the
apophyses, protrude anteriorly to form a joint with the valve
in front.

On the top of the elevations, as well as on the ribs, the
openings of the aesthetes can be seen (Fig. 4), with a
megapore (diameter= 11-14 um) in the center, surrounded by
4-9 micropores (diameter= 9 um). On the lateral areas, there
are more micropores per megapore than in the median fields.
The same is true for the absolute number of the aesthetes

American Malacological Bulletin, Vol. 6(1) (1988):153-159

153

154 AMER. MALAC. BULL. 6(1) (1988)

a

Fig. 1. Schematic cross section through an adult Lepidopleurus ca-
jetanus, left half (a, articulamentum; bc, body cavity; f, foot; g, girdle
covered with spicules; gi, gill; |, lateral fold; s, secretory cells of the
foot epithelium in the pallial groove; t, tegmentum with numerous
incorporated aesthetes) (adapted from Maile, 1981).

(number of megapores), with about 150 per mm? in the lateral
and 90 per mm2 in the median area. The head valve has the
highest density of aesthetes, about 200 per mm2.

In untreated shell valves, each megapore is filled with
the apical cap of the main stem (= megalaesthete) of an
aesthete (Fig. 2). Each micropore contains the subsidiary cap
of a micraesthete, which is a branch from the megalaesthete.
In older aesthetes, the apical caps show a perforation with
many pores of about 0.1 um in diameter (Fig. 5). The subsidiary
caps do not exhibit such a pattern. In young aesthetes the
apical cap is completely covered by the periostracum.

AESTHETES

The aesthetes are, like the papillae in the girdle, ex-
tensions of the epidermis. Most of the cells of the aesthete
are still connected with the epithelium via the aesthete canal.
Some of these basal cell extensions are nervous elements
and run further to the lateral nerve cords.

Each aesthete is about 110 nm long and 30 um thick.
It contains 35 to 40 cells of five distinct cell types: secretory
cells, central cells, photoreceptor cells, micraesthete cells
branching from the main stem, and peripheral cells (Fig. 2).
Except for the micraesthetes, every type can exhibit a basal
extension to the epithelium (Fischer, 1978a); for the
micraesthete cells the situation is not yet clear. At the shell
surface the main stem is covered by the apical cap and each
micraesthete by a subsidiary cap.

APICAL CAP. The apical cap consists only of organic
material and can be divided into two zones (Fig. 2): the distal
part, containing numerous parallel pores and the proximal
part, consisting of a network of thin filaments of two types.
There are filaments of about 80 nm in diameter, which form

10 um 2

Fig. 2. Schematic longitudinal section through an aesthete (ac, apical
cap; aec, aesthete canal; c, central cell; mi, micraesthete; n, neurites;
p, peripheral cell; ph, photoreceptor cell; sc, subsidiary cap; se,
secretory cell).

the skeleton, from which 25 nm wide filaments branch off (Fig.
6). These fine filaments form the border of the cap to the in-
terior of the aesthete.

SECRETORY CELLS. Each aesthete has three to eight tall
secretory cells of different forms. Some of them, especially
in young aesthetes, show a high metabolic activity in the pro-
ximal part; granular endoplasmic reticulum (ER), a few Golgi
apparatus and numerous mitochondria surround an active
nucleus. The secretory granules produced are stored distal-
ly. Most of the secretory cells are densely filled with mem-
brane bound secretion granules of various electron densities
(Fig. 7). The nucleus lies basally, its chromatin is highly con-
densed (Fig. 8). Remains of endoplasmic reticulum and a few
mitochrondria are often present nearby. One or two secretory
cells that open distally secrete material beneath the apical
cap. Some cytoplasm between the former granules remains;
the interior is now continuous with the extracellular space
beneath the apical cap (Fig. 9). In the neighbourhood of these

FISCHER: AESTHETES IN LEPIDOPLEURUS CAJETANUS 155

Fig. 3. Left half of an intermediate shell valve. KOH-treated (ap, apophyse; la, lateral triangle; m, median triangle). Fig. 4. Higher magnification
of the lateral triangle, KOH-treated (arrows indicate several smaller micropores surrounding a megapore). Fig. 5. Surface of an apical cap. The
cap is perforated by numerous small pores. Fig. 6. Longitudinal section of the basal area of an apical cap consisting of a network of larger

and smaller (arrows) organic filaments.

cells, some of the peripheral cells exhibit characteristics of
decomposing cytoplasm; lysosomes and autophagous
vacuoles surround an active nucleus.

CENTRAL CELLS. The central cells (‘‘sensory cells’’ ac-
cording to Boyle, 1974) (as the photoreceptor cell is also sen-
sory, | use the more neutral term ‘‘central cell’) of
Lepidopleurus cajetanus are prominent compared with the
other species studied so far. The nuclei of all central cells
(about five per aesthete) are situated in the distal part of the
aesthete. Distally, underneath the apical cap, each central cell
forms numerous microvilli and one cilium (9+2 structure) (Fig.
10). The cytoplasm of the central cells contains numerous
mitochondria and microtubules running along the long axis
of the cells. Distally the cytoplasm is filled with clear 0.3 um
wide vesicles. The central cells are connected together by
zonulae adhaerens and septate junctions.

PHOTORECEPTOR CELLS. Most of the aesthetes (but not
all of them, irrespective of the position in different valve areas)
contain one or two photoreceptor cells. They lie peripherally
in the aesthete and do not exhibit a special orientation pat-
tern, such as being always located on the same side of the

aesthete body, as it is the case in Chiton olivaceus Spengler
(Fischer and Renner, 1978). As in other species, they show
two distinct areas, the cell body and the rhabdomere (Fig.
11). The microvilli (0.05-0.1 zm in diameter) of the rhabdomere
branch from the whole distal part of the cell; they have no
regular orientation. Their cytoplasm contains small granules.
One or two cilia (9+2 structure) can be present.

The nucleus is relatively large (6.5 um) and has only
a little condensed chromatin. In the perikaryon, numerous
mitochondria, microtubules, glycogen and multivesicular
bodies are present. A specialized agranular ER forms large
areas of parallel membrane cisternae that are connected with
the granular ER. Laterally these cisternae give off numerous
clear vesicles (40-170 nm) that are found up to the rhabdome.

MICRAESTHETES. All micraesthetes branch off from the
same zone of the main stem. Their nuclei lie in this area;
they are large (6 um) and have only little condensed
chromatin. Here and in the ‘‘arm’’ (the part between the main
stem of the aesthete and the tip of the micraesthete cell) we
find numerous mitochondria and microtubules along the long
axis (Fig. 12). In the basal part, multivesicular bodies or
lysosomes are frequently found. Peripheral cells surround the

156 AMER. MALAC. BULL. 6(1) (1988)

proximal part of the ‘‘arm’’; both cells show invaginations in-
to the other cells. The ‘“‘head”’ (the tip of the micraesthete cell)
is slightly swollen and also contains mitochondria. The distal
part forms numerous microvilli towards the subsidiary cap
(Fig. 13). The ‘‘head’”’ can show a high degree of vacuoliza-
tion in some micraesthetes, but this zone does not continue
into the ‘‘arm’’.

SUBSIDIARY CAP. In contrast to the apical cap, the sub-
sidiary caps appear continuous at their outer and inner sur-
faces. They also consist of organic material. They contain in-

ner pores (width= 0.1 wm), but in nearly all cases they are
closed to the outside, as well as to the interior of the
micraesthete, by continuous sheets. The distal sheet is up to
0.2 um, the proximal sheet about 0.1 wm wide.

In some cases the micraesthete cap has been dam-
aged by organisms. In these cases, parallel sheets of vary-
ing thickness are placed underneath the remainder of the cap.
Sometimes, the subsidiary cap is completely replaced by this
structure.

PERIPHERAL CELLS. The peripheral cells surround the

Fig. 7. Longitudinal section of an aesthete nearly completely filled with the granules of secretory cells (sl, secondary lysosome). Fig. 8. Base
of a secretory cell (nu, highly condensed nucleus; sg, secretion granule). Fig. 9. Distal tip of a secretory cell after secretion. Surrounding
cytoplasm (arrows) and small cytoplasmic remains are visible between the former secretion granules. The interior is now continuous with the
extracellular material underneath the apical cap, in which microvilli and cilia (double arrow) of central cells are embedded (c, central cell).

FISCHER: AESTHETES IN LEPIDOPLEURUS CAJETANUS 157

Fig. 10. Distal tip of central cells with protruding microvilli (mv) and cilia (double arrow). Arrow indicates basal body in a central cell.

body of the aesthete as a sheet about 0.75 pm in width. They
are not present in all parts of an aesthete (e.g. Fig. 7). The
fine structure varies considerably, e.g. in the content of decom-
posing structures. In the basal part of the aesthete, peripheral
cells form an extracellular sheet of fine filaments into the shell
material. Some of the filaments: protrude, roughly perpen-
dicularly, far into the shell substance.

AESTHETE CANAL. The aesthete canal is surrounded by
peripheral cells (Fig. 14). In the center, various fibers (basal
extensions of the aesthete cells to the epithelium) run towards
the epithelium under or lateral to the shell valves. Some of
the fibers connect the secretory cells with the epithelium;
these fibers contain mitochondria and microtubules. About
ten of the fibers are much thinner than those of the secretory
cells (0.4 versus 1.5 um). They are densely filled with
microtubules and are probably nervous elements (Fig. 14).
Structures resembling neurosecretory elements are also a
regular feature in this area.

DISCUSSION

The general structure of the aesthetes of Lepidopleurus
cajetanus is very similar to that of previously studied species.
Despite the differences in the architecture of the shell valves,
there is no major difference between the aesthetes in all three
extant polyplacophoran suborders. The aesthetes are obvious-

ly an evolutionarily old system in the chitons.

The different cell types, each with pronounced ultra-
structural characteristics, suggest that the aesthetes are com-
pound organs with both a sensory and a secretory function.
It is not clear whether some or all cell types work together
to perform a more complex function or whether they function
more or less independently.

The secretory cells produce secretions basally and
release them apically. In Chiton olivaceus, animals outside
the water show an increased secretory activity (Fischer,
1978a). Additionally, recordings with a glass microelectrode
show slow rhythmic changes in the electrical potential under
the same conditions (Fischer, unpub. data). This could sug-
gest that one function of the secretion is to prevent the desi-
ccation of the aesthetes during low tide. However, species that
prefer to live in deeper water also have well developed
secretory cells (Baxter et a/., 1987). The secretions probably
have other protective functions, e.g. against predators or
organisms growing on the shell. One indication for such a
function is that the pores of the apical cap open only in older
aesthetes, whereas newly formed aesthetes are covered by
the periostracum.

The role of the central cells is not known. In their fine
structure they resemble chemoreceptors of insects (Ernst,
1969). The position of this cell type underneath the perforated
apical cap, the pronounced membrane proliferations (microvilli
and cilia), and the high metabolic activity support such a
hypothesis. Structures resembling nervous elements were

158 AMER. MALAC. BULL. 6(1) (1988)

Fig. 11. Photoreceptor cell at the transition of perikaryon and rhabdomere (mvb, multivesicular body; nu, nucleus; rh, rhabdomere; ser, specialised
agranular ER). Fig. 12. Longitudinal section of the ‘‘arm’’ part of a micraesthete cell (mit, mitochondrion; mt, microtubules). Fig. 13. Sub-
sidiary cap (sc) with the microvilli (mv) of the underlying micraesthete cell. Fig. 14. Longitudinal section through an aesthete canal (nu, nucleus

of a peripheral cell). Arrows indicate profiles resembling neurites.

found near the base, but their relationship to the central cells
is not clear. A possible function could be to detect desicca-
tion and/or animals grazing on the shell (and eating also the
distal parts of the aesthetes), with a subsequent reaction of
the secretory cells. Lepidopleurus cajetanus has the best
developed central cells of all species studied so far; in the
Lepidopleurina the articulamentum is lacking in broad shell
areas and the aesthetes are connected directly with the dor-
sal epithelium. Destruction of the aesthetes and a subsequent
invasion of microorganisms into empty aesthete canals could
be much more severe in this group than in the Ischnochitonina
or the Acanthochitonina.

The photoreceptor cells resemble in detail the photo-
receptor cells of Chiton olivaceus and Acanthochitona
fascicularis (Fischer, 1978b, 1979; Fischer and Renner, 1978)
as well as the photoreceptor cells in the two different shell-
eye types in the chitons (Boyle, 1969; Haas and Kriesten,
1978). Boyle (1974) found no typical photoreceptor cells in the
aesthetes of Lepidochitona cinerea L., but he described
“microvillous areas’ and areas with ‘‘lamellate bodies’. These
very likely correspond to the rhabdomere and the agranular
ER of the photoreceptor cells. This cell type is the only one

in the aesthete that shows ultrastructural differences be-
tween light- and dark-adapted animals (Fischer, 1978a). Ad-
ditionally, experiments with partially masked C. olivaceus
clearly show that the shell valves contain photoreceptive
elements (Fischer, unpub. data). As a common feature,
photoreceptor cells seem to be a primary part in chiton
aesthetes.

At first sight, it is astonishing that the shell valves con-
tain so many, and simple, photoreceptive elements. In some
species, aesthetes in certain shell areas have been transform-
ed into eyes of various complexity (Boyle, 1969; Fischer, 1978a;
Haas and Kriesten, 1978). Most species, however, have only
the ‘‘normal’’ aesthete type. The situation could be com-
parable with other invertebrates, like the earthworm which
avoid light during the day and feed at dawn and when it is
dark. The earthworm also has many primitive light receptors
dispersed in the skin. As behavioural studies show (Fischer,
unpub. data), the photoreceptor cells in the aesthetes have
a similar function. Most chiton species avoid bright sunlight
and hide below stones or in the mud during the day. Chitons
with masked shells do not exhibit such a behaviour (except
species that also have photoreceptor cells in their girdle

FISCHER: AESTHETES IN LEPIDOPLEURUS CAJETANUS 159

papillae, e.g. Acanthochitona fascicularis). Chitons that live
in deeper water obviously have lost their photoreceptor cells.
Baxter et a/. (1987) found no photoreceptor cells in the
aesthetes of Jonicella marmorea Fabricius.

The function of the micraesthetes is still completely
obscure. Their high density in the shell valves suggests that
they play an important role in the biology of the Poly-
placophora. Among all species studied, only some of the
lateral aesthetes in Acanthochitona fascicularis lack
micraesthetes (Fischer, 1979). Baxter et a/. (1987) showed that
in Tonicella marmorea, the micraesthetes contain numerous
lamellate granules. They suggest that micraesthetes and the
megalaesthete produce periostracum material. This
hypothesis has already been put forward by Nowikoff (1907).
In all the species | studied, no secretory granules in the
micraesthetes could be found. In addition, the subsidiary cap
does not allow a penetration of material from the inside of the
aesthete to the outside. In electrical recordings from the area
underneath the subsidiary cap in Chiton olivaceus, no indica-
tion of a secretory process could be found under many dif-
ferent experimental conditions (Fischer, unpub. data). In other
species, the subsidiary cap looks similar to that of
Lepidopleurus cajetanus (Fischer and Renner, 1978; Haas and
Kriesten, 1978) or are much thinner but without inner pores
(Boyle, 1974). Both types can be present in different shell areas
in certain species (Fischer, 1979). Baxter et al. (1987) showed
that in 7. marmorea, microvilli of the micraesthete cell pro-
trude into the subsidiary cap. However, the distal surface of
the cap also seems to be continuous in this species. In all
species studied so far, the continuous part of the subsidiary
cap is about 0.4 um, irrespective to whether inner pores are
present. Certainly there is a great need to study the aesthetes
of species that differ in their ecology from the species studied
so far, in order to gain a better understanding of the function
of the micraesthetes.

ACKNOWLEDGMENTS

| am most grateful to Prof. G. A. Manley for correcting my

English. | thank Birgit Seibel for technical assistance and Renate Kam-
merer for reading the manuscript. The present paper arose from a
talk presented at the Symposium on the Biology of the Polyplacophora
at the 1987 meeting of the American Malacological Union. | thank
the AMU for financial support to attend that meeting.

LITERATURE CITED

Baxter, J. M., A. M. Jones and M. G. Sturrock. 1987. The ultrastruc-
ture of aesthetes in Tonicella marmorea (Polyplacophora;
Ischnochitonina) and a new functional hypothesis. Journal of
Zoology (London) 211:589-604.

Boyle, P. R. 1969. Fine structure of the eyes of Onithochiton neglec-
tus (Mollusca: Polyplacophora). Zeitschrift fur Zellforschung
102:313-332.

Boyle, PR. R. 1974. The aesthetes of chitons. 2. Fine structure in
Lepidochitona cinereus. Cell Tissue Research 153:383-398.

Ernst, K. D. 1969. Die Feinstruktur der Riechsensillen auf der Anten-
nen des Aaskafers Necrophorus. Zeitschrift fur Zellforschung
94:72-102.

Fischer, F. P. 1978a. Untersuchungen an den Astheten dreier Poly-
placophoren-Arten. Doctoral Dissertation. Ludwig-Maximilians-
Universitat Munchen. 118 pp.

Fischer, F. P. 1978b. Photoreceptor cells in chiton aesthetes (Mollusca,
Polyplacophora). Spixiana 1:209-213.

Fischer, F. P. 1979. Die Astheten von Acanthochitona fascicularis
(Mollusca, Polyplacophora). Zoomorphologie 92:95-106.

Fischer, F. P. and M. Renner. 1978. Die Feinstruktur der Astheten von
Chiton olivaceus (Mollusca, Polyplacophora.) He/lgolander
wissenschaftliche Meeresuntersuchungen 31:425-443.

Haas, W. and K. Kriesten. 1978. Die Astheten mit intrapigmentarem
Schalenauge von Chiton marmoratus L. (Mollusca, Placo-
phora). Zoomorphologie 90:253-268.

Maile, W. 1981. Drusenzellen und Drusenzellenkomplexe an Mantel,
Fuss and Kiemenrinne dreier Polyplacophoren-Arten. Diplo-
marbeit, Ludwig-Maximilians-Universitat Munchen. 136 pp.

Nowikoff, M. 1907. Uber die Ruckensinnesorgane der Placophoren
nebst einigen Bemerkungen uber die Schale derselben.
Zeitschrift fur wissenschaftliche Zoologie 88:154-186.

Reynolds, F. S. 1963. The use of lead citrate at high pH as an elec-
tron opaque stain in electron microscropy. Journal of Cell
Biology 17:208-212.

Date of manuscript acceptance: 16 October 1987

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161

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162

Respectfully submitted,
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54th ANNUAL MEETING
THE AMERICAN MALACOLOGICAL UNION
CHARLESTON, SOUTH CAROLINA
RADISSON FRANCIS MARION HOTEL
JUNE 19 - 26, 1988

The 54th annual meeting of the American Malacological Union will be held June
19 - 26, 1988, in Charleston, South Carolina. Charleston is a historical city, many parts
of which have been beautifully restored as has the Radisson Francis Marion Hotel
which is located downtown within walking distance of many restaurants, shops and
other attractions. Charleston is easily accessible both by air and by interstate highway.

Three symposia are planned:

APPLICATIONS OF NUCLEIC ACID TECHNIQUES TO MOLLUSCAN SYSTEMATICS
(Organized by Dr. M. G. Harasewych, Dept. of Invertebrate Zoology,
Smithsonian Institution)

SYSTEMATICS AND EVOLUTION OF NON-MARINE MOLLUSKS
(Organized by Dr. Robert Hershler, Dept. of Invertebrate Zoology,
Smithsonian Institution)

HISTORY OF MALACOLOGY
(Organized by Dr. W. Backhuys, Leiden, The Netherlands)

In addition to the symposia, contributed papers and poster presentations, scheduled events will in-
clude a tour of historic Charleston, guided field trips to terrestrial and marine molluscan communities,
an auction to benefit the symposium fund, and a banquet.

For further information please contact:
Richard E. Petit
President, AMU
P. O. Box 30
North Myrtle Beach, South Carolina 29582, USA

163

Western Society of Malacologists

ANNOUNCEMENT AND INVITATION TO PARTICIPATE

Symposium on Biogeography and Evolution of the Molluscan
Fauna of the Galapagos Islands

21st Annual Meeting of the Western Society of Malacologists
Sonoma State University, Rohnert Park, California

17-21 July 1988

The Western Society of Malacologists maintains a long-standing tradition of emphasis on eastern Pacific
molluscan faunas and in keeping with this tradition a symposium will be held on Monday, July 18, 1988 in
Darwin Hall on the campus of Sonoma State University on the biogeography and evolution of the molluscan
fauna of the Galapagos Islands. The purpose of this symposium is to bring together, some 150 years after
Charles Darwin visited the Galapagos, researchers with interests in the taxonomic composition, biogeographic
affinities, and evolutionary history of the living and fossil molluscan fauna of the Galapagos.

The following is a preliminary list of symposium participants; additional contributors are being solicited:

J. Wyatt Durham - University of California, Berkeley

William K. Emerson - American Museum of Natural History, New York
Terrence M. Gosliner - California Academy of Sciences, San Francisco
Michel Montoya - San Jose, Costa Rica

Carole S. Hickman - University of California, Berkeley

Matthew J. James - Sonoma State University

Shi-Kuei Wu - University of Colorado, Boulder

Donald R. Shasky - Redlands, California

William D. Pitt - Sacramento, California

David K. Mulliner - San Diego, California

E. Alison Kay - University of Hawaii, Manoa

Victor A. Zullo - University of North Carolina, Wilmington

Jere H. Lipps - University of California, Davis

Contributions are welcome from neontologists and paleontologists with an interest in any aspect of
taxonomic composition, biogeographic affinities, evolutionary relationships, stratigraphic distribution, geologic
context, oceanographic setting or paleoecological relationships.

For further information, please contact:

Dr. Matthew J. James

WSM - Galapagos Symposium
Department of Geology

Sonoma State University

Rohnert Park, California 94928, U. S. A.
Phone: (707) 664-2301 or 2334

164

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Vail, V. A. 1977. Comparative reproductive anatomy

“ae age of 3 viviparid gastropods. Malacologia

1 ae 16(2):519-540. )

ae Yonge, C. M. and T. E. Thompson. 1976. Living

aan Marine Molluscs. William Collins Sons & Co.,
Ltd., London. 288 pp.

Beattie, J. H., K. K. Chew, and W. K. Hershberger.
1980. Differential survival of selected strains of
Pacific oysters (Crassostrea gigas) during
summer mortality. Proceedings of the National

tt Shellfisheries Association 70(2):184-189.

POR Seed, R. 1980. Shell growth and form in the Bivalvia.

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\

S) AMERICAN
* MALACOLOGICAL
\; BULLETIN

VOLUME 6 NUMBER 2 OCTOBER 1988

CONTENTS

A comparative study of late prehistoric and modern molluscan faunas
of the Little Pigeon River System, Tennessee. PAUL W. PARMALEE..................... 165

Evaluation of techniques for age determination of freshwater mussels (Unionidae).
- RICHARD J. NEVES and STEVEN N. MOYER.................... 00.0000 cece eee 179

intracapsular development of Thais haemastoma canaliculata (Gray) (Prosobranchia:
Muricidae) under laboratory conditions. RICHARD A. ROLLER and
ele SESS CST Se Te a ee Siege oa IN ch eer a Co Oe ic a 189

Temporal and spatial variation of shell microstructure of Polymesoda caroliniana pez
(Bivalvia: Heterodonta). ANTONIETO TAN TIU.............. 000... 00... e eee eee 199

The use of arm sucker number in octopodid systematics (Cephalopoda:
Pecusmocley arto MALES: TOR shin fe ka ei din Dee ee te epee a els vies ghee 207

Research Note: Effects of fixation and preservation methods on the morphology
of a loliginid squid (Cephalopoda: Myopsida). JOSE MILTON

ANDRIGUETTO, JR. and MANUEL HAIMOVIC!.............. 00... 0.00. eee 213
ee 2 DSS Dog greet » CE RDRINS bee ele See ee eared OAS Ne 219
CAWUIMVETP LUNG. G A Se SS ic Be Rae A, og DSO er a ON 6) ESE Meir im UN: ADO RR SO Cet 2 aed 220
Se MeMMETII CANNER Le rae NR eS at eo Fe A RSs he Soe Phe BR ee his Bee alibaat ei aee gt ey 223
RECN NA NITRA RCM etch te PNR se lg see dS RG eS ea houhe Wb fee CN bun Wien ate nc dee be he Soedeone Ba 254

Seeea NG OME ee co POR Sc ce ek eS its ae sis eek ake Bs anh opie WORRY Papbeweied of: 285

AMERICAN MALACOLOGICAL BULLETIN

EDITOR-IN-CHIEF

ROBERT S. PREZANT
Department of Biology
Indiana University of Pennsylvania
Indiana, Pennsylvania 15705

MELBOURNE R. CARRIKER
College of Marine Studies
University of Delaware
Lewes, Delaware 19958

GEORGE M. DAVIS

Department of Malacology

The Academy of Natural Sciences
Philadelphia, Pennsylvania 19103

R. TUCKER ABBOTT
American Malacologists, Inc.
Melbourne, Florida, U.S.A.

JOHN A. ALLEN
Marine Biological Station
Millport, United Kingdom

JOHN M. ARNOLD
University of Hawaii
Honolulu, Hawaii, U.S.A.

JOSEPH C. BRITTON
Texas Christian University
Fort Worth, Texas, U.S.A.

JOHN B. BURCH
University of Michigan
Ann Arbor, Michigan, U.S.A.

EDWIN W. CAKE, JR.
Gulf Coast Research Laboratory
Ocean Springs, Mississippi, U.S.A.

PETER CALOW
University of Sheffield
Sheffield, United Kingdom

BOARD OF EDITORS

ASSOCIATE EDITORS

MANAGING EDITOR

RONALD B. TOLL
Department of Biology
University of the South

Sewanee, Tennessee 37375

W. D. RUSSELL-HUNTER
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RICHARD E. PETIT
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North Myrtle Beach, South Carolina 29582

BOARD OF REVIEWERS

JOSEPH G. CARTER
University of North Carolina
Chapel Hill, North Carolina, U.S.A.

ARTHUR H. CLARKE
Ecosearch, Inc.
Portland, Texas, U.S.A.

CLEMENT L. COUNTS, lil
Coastal Ecology Research
University of Maryland

Princess Anne, Maryland, U.S.A.

THOMAS DIETZ
Louisiana State University
Baton Rouge, Louisiana, U.S.A.

WILLIAM K. EMERSON
American Museum of Natural History
New York, New York, U.S.A.

DOROTHEA FRANZEN
Iilinois Wesleyan University
Bloomington, Illinois, U.S.A.

VERA FRETTER

University of Reading
Berkshire, United Kingdom

ISSN 0740-2783

Syracuse, New York 13210

THOMAS R. WALLER

Department of Paleobiology
Smithsonian Institution
Washington, D. C. 20560

ROGER HANLON
University of Texas
Galveston, Texas, U.S.A.

JOSEPH HELLER
Hebrew University of Jerusalem
Jerusalem, Israel

ROBERT E.'HILLMAN
Battelle, New England
Duxbury, Massachusetts, U.S.A.

kK. ELAINE HOAGLAND
Association of Systematics Collections
Washington, D.C., U.S.A.

RICHARD S. HOUBRICK
U.S. National Museum
Washington, D.C., U.S.A.

VICTOR S. KENNEDY
University of Maryland
Cambridge, Maryland, U.S.A.

ALAN J. KOHN
University of Washington
Seattle, Washington, U.S.A.

LOUISE RUSSERT KRAEMER
University of Arkansas
Fayetteville, Arkansas, U.S.A.

JOHN N. KRAEUTER
Baltimore Gas and Electric
Baltimore, Maryland, U.S.A.

ALAN M. KUZIRIAN

NINCDS-NIH at the

Marine Biological Laboratory
Woods Hole, Massachusetts, U.S.A.

RICHARD A. LUTZ
Rutgers University
Piscataway, New Jersey, U.S.A.

EMILE A. MALEK
Tulane University
New Orleans, Louisiana, U.S.A.

MICHAEL MAZURKIEWICZ
University of Southern Maine
Portland, Maine, U.S.A.

JAMES H. McLEAN
Los Angeles County Museum
Los Angeles, California, U.S.A.

ROBERT F. MCMAHON
University of Texas
Arlington, Texas, U.S.A.

ROBERT W. MENZEL
Florida State University
Tallahassee, Florida, U.S.A.

ANDREW C. MILLER
Waterways Experiment Station
Vicksburg, Mississippi, U.S.A.

BRIAN MORTON
University of Hong Kong
Hong Kong

JAMES J. MURRAY, JR.
University of Virginia
Charlottesville, Virginia, U.S.A.

RICHARD NEVES

Virginia Polytechnic Institute
and State University

Blacksburg, Virginia, U.S.A.

JAMES W. NYBAKKEN
Moss Landing Marine Laboratories

Moss Landing, California 95039-0223

WINSTON F. PONDER
Australian Museum
Sydney, Australia

CLYDE F. E. ROPER
U.S. National Museum
Washington, D.C., U.S.A.

NORMAN W. RUNHAM
University College of North Wales
Bangor, United Kingdom

AMELIE SCHELTEMA

Woods Hole Oceanographic Institution

Woods Hole, Massachusetts, U.S.A.

ALAN SOLEM
Field Museum of Natural History
Chicago, Illinois, U.S.A.

DAVID H. STANSBERY
Ohio State University
Columbus, Ohio, U.S.A.

FRED G. THOMPSON
University of Florida
Gainesville, Florida, U.S.A.

THOMAS E. THOMPSON
University of Bristol
Bristol, United Kingdom

NORMITSU WATABE
University of South Carolina
Columbia, South Carolina, U.S.A.

KARL M. WILBUR
Duke University
Durham, North Carolina, U.S.A.

Cover. The intracapsular development of the muricid Thais haemastoma canaliculata is discussed in an article by Roller and Stickle in this

volume, pages 189-197.

THE AMERICAN MALACOLOGICAL BULLETIN is the official journal publication of the American Malacological Union.

AMER. MALAC. BULL. 6(2)
October 1988

A COMPARATIVE STUDY OF LATE PREHISTORIC
AND MODERN MOLLUSCAN FAUNAS OF THE
LITTLE PIGEON RIVER SYSTEM, TENNESSEE

PAUL W. PARMALEE
FRANK H. McCLUNG MUSEUM
UNIVERSITY OF TENNESSEE
KNOXVILLE, TENNESSEE 37996, U. S. A.

ABSTRACT

Shells of freshwater gastropods and naiads recovered during the period June - December 1985
at the McMahan site, an aboriginal Mississippian (Dallas component: AD 1300-1600) mound and village
complex situated adjacent to the West Prong Little Pigeon River, Sevierville, Sevier County, Tennessee
comprised the largest prehistoric molluscan species assemblage from a small river in East Tennessee
yet known. Six species of aquatic gastropods (7,411 shells) and 3,855 valves of freshwater mussels
(Bivalvia: Unionidae), representing 45 species, were identified. Three of the six species of gastropods
and 31 of the 45 species of mussels no longer occur in the Little Pigeon River system. For a 24 month
period, June 1985 - May 1987, extant mussel populations in the West Prong Little Pigeon River adja-
cent to the McMahan site were monitored and shells collected, primarily from muskrat feeding sta-
tions. Only 11 species occur as viable populations; urbanization with its accompanying pollution prob-
ably represents the major cause in decimating the rich molluscan assemblage present during the late

prehistoric period.

The McMahan site (40SV1), a multi-component, late
prehistoric aboriginal village and mound complex situated ad-
jacent to the West Prong Little Pigeon River, now within the
city limits of Sevierville, Sevier County, Tennessee has
aroused the interest of both amateur and professional arch-
aeologists for over a century. The mound, Late Mississippian
(Dallas component: AD 1300-1600) in origin, was reported to
have been 125 yards (112 m) from the river and was 16 feet
(4.8 m) in height and 240 feet (72 m) in circumference at the
time Edward Palmer ‘“‘opened”’ it in September, 1881 (Holmes,
1884). Palmer, then employed by the Bureau of Ethnology,
recovered numerous lithic artifacts, ceramic vessels, engraved
marine shell gorgets and three species of marine gastropods
(listed as ‘‘Marginella?, Oliva?, Busycon perversum’’) that had
been fashioned into beads and other objects. These were
found as burial accouterments. Also listed in the 1884 report
were three species of freshwater gastropods and four species
of naiads.

Approximately 50 years passed before the mound was
again excavated, this time by George Barnes, a relic collec-
tor from Tennessee who, like Palmer, removed numerous
burials and quantities of lithic, ceramic and shell artifacts en-
countered in association with them. Except for surface col-
lecting, little attention was given to the surrounding village

areas until June - August 1978 when highway (TN Rt. 441 N
Bypass) salvage excavations were carried out by Dr. Brian
Butler for the Tennessee State Division of Archaeology. A
series of test pits in the area to be affected by highway con-
struction, ca. 1,500 m south of the mound, revealed former
occupation of the site by Middle Woodland (Connestee: AD
300-600), Mississippian (Dallas: AD 1300-1600), and Cherokee
(ca. AD 1650-1800) peoples. Bone from the various excava-
tion units was generally well preserved, but shell was not. For
this reason, and particularly because the majority of faunal
material recovered came from pits and various other features
that contained a mixture of Connestee, Dallas and/or
Cherokee cultural materials, shell identifications and counts
from these excavations were not incorporated in this study.
It snould be noted, however, no species were recovered in
Butler’s excavations that were not represented in the mound
and adjacent village areas occupied by Dallas inhabitants.

METHODS

Owner of the property that included the mound and
remaining former village areas of the McMahan site, Mr.
James A. Temple of Sevierville arranged for the removal and
sale of the site (but preserving most of the mound) for topsoil

American Malacological Bulletin, Vol. 6(2) (1988):165-178

165

166 AMER. MALAC. BULL. 6(2) (1988)

in the early 1980s. By the end of 1983, the soil on the north
side of the mound had been removed and stockpiled. It was
not possible to undertake salvage operations at that time, so
only a small sample of bone and shell was recovered
periodically from the stockpiles as they were removed over
a period of months. In order to determine the perimeter of
the mound along its south-facing edge so as not to destroy
that portion of it during soil removal, Mr. Richard R. Polhemus,
Research Associate, Frank H. McClung Museum, University
of Tennessee, at the owner’s request excavated a north - south
trench (0.5 m wide, 21 m long, and 1.2 - 2.0 m deep from about
mid-point to the south edge) to determine stages of construc-
tion and location of its outermost edge. Preservation of bone
and shell from the mound fill was generally good to excellent;
since the mound was built by Late Mississippian (Dallas)
peoples and was part of the adjoining village complex from
which the majority of faunal materials were recovered, shell
from the trenching excavation was combined with the village
material for this analysis.

Removal of the remaining village area south of the
mound began in May 1985 and was completed by December
of that year. Funds could not be made available for an organ-
ized archaeological salvage operation, so the only alternative,
if any data were to be obtained from the wealth of both cultural
and faunal materials present, was to recover as much as
possible in the allotted time by the author’s personal effort.
| visited the site on 34 days during the period of soil removal,
averaging ca. five hours each visit. On six occasions
volunteers provided assistance with the excavation of material
which was accomplished for the most part by shovelling and
trowel sorting. The area was surface collected on each visit
and the growing stockpiles of topsoil were also searched for
cultural and faunal remains. Days in which soil removal was
in progress, each newly exposed feature resulting from cuts
(profiles) made by the heavy equipment was carefully exam-
ined for its content. Shell recovered from two five-foot test
squares excavated at the south edge of the mound in October
1985 by Richard Polhemus was incorporated with those from
the village excavations, surface collections, and mound. All
recovered cultural and faunal specimens were washed and
cleaned with a soft brush; after drying each collection lot was
labelled and eventually a large series of the identifiable shells
was also given the site designation number and date
recovered. All specimens have been incorporated into the
Frank H. McClung Museum collections, The University of Ten-
nessee, Knoxville.

Recognizing the species diversity present in the ar-
chaeological molluscan samples from the McMahan site, a
study of gastropod and freshwater mussel populations
presently inhabiting the Little Pigeon River system was under-
taken to determine possible changes in extant assemblages
compared with those that existed in late prehistoric times. The
Little Pigeon River is fed by two major tributaries, the East
Fork, a small second order stream, and the West Prong Lit-
tle Pigeon River, a fifth order stream only slightly less in size
than the Little Pigeon itself (Fig. 1). Although the East Fork
and the Little Pigeon were collected periodically, survey and
collecting emphasis was placed on a ca. 0.7 km stretch of

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Fig. 1. Map showing the Little Pigeon River system and location of
the McMahan site.

the West Prong Little Pigeon River that flowed above, adja-
cent to and below the McMahan site. Collecting trips were
made in this section of the river at least twice each month
for a 24-month period beginning in June 1985 and ending in
May 1987. A total of 54 collecting and survey trips were made
in this section of river during this period. Muskrats (Ondatra
zibethica Linnaeus, 1766) inhabit the banks of the river and
are the major predator of bivalves; utilization of this food
resource is greatest during the winter months, ca. November
through March. Shells obtained from muskrat feeding stations
and those scattered along the river bottom, also probably
discarded after the animal had been eaten by muskrats,
formed the basis on which an evaluation of species occur-
rence and population density was made. Notations were made
of live individuals and their number when encountered, but
with the exception of less than a dozen specimens no living
naiads were collected. Voucher specimens of most species
represented have been placed in the Department of
Malacology, Academy of Natural Sciences of Philadelphia,
Philadelphia, Pennsylvania and the Museum of Zoology, The
Ohio State University, Columbus, Ohio; most of the remain-
ing specimens obtained during this study are housed in the
Malacology Collection, Frank H. McClung Museum.

RESULTS

SPECIES ACCOUNTS: GASTROPODA

Shells of six species of aquatic gastropods were
recovered at the McMahan site (Table 1); 93% of the 7,411

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS 167

Table 1. Freshwater gastropod shells identified from the Dallas com-
ponent, McMahan site (40SV1), Sevierville, Sevier County, TN.

No. of % of

Species Shells Shells
Campeloma cf. decisum (Say, 1816) 38 51
lo fluvialis (Say, 1825) 374 5.05
Leptoxis praerosa (Say, 1821) 3,860 52.08
Lithasia (Angitrema) verrucosa
(Rafinesque, 1820) 10 13
Pleurocera canaliculatum (Say, 1821) 94 1.27
P parvum (Lea, 1862) 3,035 40.95
Totals 7,411 99.99

specimens identified were those of Pleurocera parvum (Lea,
1862) and Leptoxis praerosa (Say, 1821), shells of the latter
species representing over half of all the aquatic gastropods
from the site. Most specimens of Leptoxis compared well with
L. praerosa, many reaching a very large size characteristic
of big river forms. Shell length (tip of the apex to the tip of
the anterior aperture canal) of 20 of the largest specimens
recovered had a mean of 18.4 mm. Although numerous small
specimens of Leptoxis appeared intermediate between L.
praerosa and the small river species L. subglobosa (Say, 1825)
in shell characteristics, they could simply reflect juvenile
stages of the former.

Specimens of the Spiny River Snail /o fluvialis (Say,
1825), comprised 5.0% of the aquatic snails. The taxonomy
of this unique species, once widespread in the upper Ten-
nessee River system, has been of special interest to
malacologists for nearly 100 years. Adams (1915) provided the
most definitive work on this gastropod up to that time; he
recorded 14 species, characterized in part on shell size and
obesity but especially on variation in spinosity. Generally, the
small river species (forms) lacked spines while those popula-
tions established in big river shoals exhibited maximum
development of spine size. Three distinct forms of /. fluvialis
occurred at the McMahan site, and Parmalee and Bogan
(1987) have discussed their taxonomy and ecological implica-
tions. Thirty-two percent lacked spines (Small river form), 47%
possessed low spines only on the last shoulder whorl (‘‘in-
termediate’ form) and 21% had well developed spines (big
river form). It can be concluded that the West Prong Little
Pigeon River possessed a varied substrate, shallow riffles and
deep shoals within a 1.6-3.2 km stretch of the site that allowed
the establishment of varied forms of /o.

Combined, shells of the three remaining species of
gastropods represented at the McMahan site comprised < 2%
of the total. Although somewhat variable in habitat preference,
Pleurocera canaliculatum (Say, 1821) and Campeloma cf.
decisum (Say, 1816) can be found most often partially buried
in mud or under matts of vegetation or debris near the shore.
Although Lithasia (Angitrema) verrucosa (Rafinesque, 1820)
can also occur in similar habitats, it apparently prefers rocks
and submerged logs in stretches of river with pronounced cur-
rent. Possibly they were less visible to the Indians while
gathering mollusks than other species that inhabit more ex-

posed river substratum. However, probable pristine river con-
ditions at that time did not include a mud or silt substratum
favorable to these species and therefore they were relatively
uncommon to rare. Judging from the size range and numbers
of gastropod shells recovered, occupants of the McMahan site
gathered whatever was available.

SPECIES ACCOUNTS: PELECYPODA

The number of naiad species represented in the
molluscan assemblage from the McMahan site relative to the
quantity of valves recovered and period of accumulation is
unequaled among other archaeologically derived samples
from Tennessee. A total of 3,855 valves, representing a
minimum of 45 species (Table 2), was identified to the generic
and/or species level. Forty-three species of freshwater mussels
were identified from the Clinch River Breeder Reactor Plant
site, Roane County (Parmalee and Bogan, 1986), but this in-
volved a sample of ca. 23,900 valves and a time span of ac-
cumulation of at !east 1,500 years. Parmalee et al. (1982)
recorded 45 species of naiads from 15 aboriginal sites in the
Chickamauga Reservoir (Tennessee River), based on the
identification of nearly 27,900 valves, but again this involved
approximately a 1,500 year time period. Nearly 3,800 valves,
representing 38 species of naiads, were recorded by Bogan
(1980) from Dallas and Cherokee occupational zones at the
Toqua site, Little Tennessee River, Monroe County. The diverse
naiad assemblage reflected in the McMahan site molluscan
sample is indicative of the rich late prehistoric populations
that inhabited this small river and provides some evidence
of the varied aquatic habitats that apparently once existed in
the West Prong Little Pigeon River.

Amblema plicata (Say, 1817): Parmalee and Bogan
(1986) noted that the Three-ridge Mussel possibly could not
have been as numerous in prehistoric times as it is at pre-
sent, judging by the relatively small numbers (2.19% of ca.
23,900 valves) recovered at the Clinch River Breeder Reac-
tor Plant site. It accounted for < 1% of 27,875 valves identified
from 15 sites in the Chickamauga Reservoir (Parmalee et ai.,
1982). Although valves of both juveniles and adults were noted
in the naiad sample from the McMahan site, their number ac-
counted for <1% of the total.

Fusconaia Simpson, 1900: Valves of both forms of F
barnesiana (Lea, 1838), the Tennessee Pigtoe F barnesiana
tumescens (Lea, 1845), a heavy, swollen shell, and F. barne-
siana bigbyensis (Lea, 1841), a thinner, more compressed form
occurred in the McMahan site samples. Ortmann (1918) noted
that ‘“..we have the phenomenon that flat and compressed
forms are found in the headwaters, swollen forms in the larger
rivers, with the intergrades between them in rivers of medium
size.’ Ortmann (1918) reported both forms from the Little
Pigeon River; combined, shells of both forms and “‘in-
tergrades’’ totalled 347, representing 9.0% of the sample.

Nearly 11% of all identified valves were those of the
Long Solid Fusconaia subrotunda (Lea, 1831), and the
number of shells (409) of this species in the McMahan site
sample ranked second in the total assemblage. At least two
distinct forms were present, one of which Ortmann (1918)

168 AMER. MALAC. BULL. 6(2) (1988)

Table 2. Freshwater mussels identified from the Dallas component,
McMahan site (40SV1), Sevierville, Sevier County, TN. [I = Interior
Basin (Mississippi); C = Cumberlandian; U = Unknown].

No. of Region of
Species Valves % — Origin

Amblema plicata (Say, 1817) 22 57 |
Fusconaia barnesiana (Lea, 1838) 347 9.00 Cc
F. subrotunda (Lea, 1831) 409 10.60 U
Quaarula cylindrica (Say, 1817) 6 15 U
Q. pustulosa (Lea, 1831) 5 12 |
Q. sparsa (Lea, 1841) 50 1.29 Cc
Cyclonaias tuberculata

(Rafinesque, 1820) 74 1.91 |
Elliptio crassidens (Lamarck, 1819) 23 59 |
E. dilatata (Rafinesque, 1820) 70 1.81 U
Hemistena lata (Rafinesque, 1820) 1 02 C
Lexington dolabelloides (Lea, 1840) 96 2.49 Cc
Plethobasus cooperianus (Lea, 1834) 11 .28 I
P. cyphyus (Rafinesque, 1820) 46 1.19 |?
Pleurobema cordatum

(Rafinesque, 1820) 33 85 |
P. oviforme (Conrad, 1834) 24 62 Cc
P. plenum (Lea, 1840) 21 54 |
P cf. rubrum (Rafinesque, 1820) 1 02 |
Alasmidonta marginata (Say, 1819) 1 02 |
A. viridis (Rafinesque, 1820) 31 80 |
Anoaonta, A. cf. grandis (Say, 1829) 1 02 |
Lasmigona costata (Rafinesque, 1820) 8 .20 U
L. holstonia (Lea, 1831) 5 12 Cc
Actinonais ligamentina (Lamarck, 1819) 148 3.83 |
Toxolasma lividus (Rafinesque, 1831) 131 3.39 Cc
Epioblasma arcaeformis (Lea, 1831) 26 67 Cc
E. brevidens (Lea, 1834) 1 02 Cc
E. capsaeformis (Lea, 1834) 42 1.08 C
E. cf. florentina (Lea, 1857) 1 02 Cc
E. haysiana (Lea, 1833) 23 59 Cc
E. stewardsoni (Lea, 1852) 2 05 Cc
E. torulosa (Rafinesque, 1820) 11 .28 C
Lampsilis fasciola (Rafinesque, 1820) 385 9.98 |
L. ovata (Say, 1817) 79 2.04 I
Lemiox rimosus (Rafinesque, 1831) 8 .20 C
Ligumia recta (Lamarck, 1819) 2 05 U
Medionidus conradicus (Lea, 1834) 172 4.46 Cc
Obovaria subrotunda (Rafinesque, 1820) 9 .23 |
Potamilus alatus (Say, 1817) 9 .23 |
Villosa iris (Lea, 1830) 167 4.33 Cc
V. trabilis (Conrad, 1834) 183 4.74 Cc
V. vanuxemensis (Lea, 1838) 302 783 Cc
V. spp. 200 5.18 —
Cyprogenia stegaria (Rafinesque, 1820) 5 12 U
Dromus dromas (Lea, 1834) 32 83 Cc
Ptychobranchus fasciolaris

(Rafinesque, 1820) 124 3.21 U
P. subtentum (Say, 1825) 508 13.17 Cc

Totals 3855 99.74

recorded as F. pilaris (Lea, 1840) and viewed it as “‘...the up-
per Tennessee representative of F subrotunda Lea of the Ohio
drainage, and it could be merely a dwarfed, globular form of
the latter?’ Apparently this form, which dominated the
McMahan site F. subrotunda ‘‘complex,’ was typical of the

large river such as the Tennessee and the lower Little Ten-
nessee and French Broad. A few valves of the compressed
headwaters form of this species were recovered. The Long
Solid appears to have been a major component of the West
Prong Little Pigeon River prehistoric naiad fauna and the
predominance of the thick globular form suggests stretches
of large river habitat.

Quaadrula Rafinesque, 1820: Three species belonging
to this genus were represented in the McMahan site naiad
assemblage; however, only six valves of the Rabbit’s Foot
Quaarula cylindrica (Say, 1817) and five valves of the
Pimpleback Q. pustulosa (Lea, 1831) were recovered. At pre-
sent both can be found locally common in small to large river
habitats throughout the state, but it has been noted (Parmalee
et al., 1982; Parmalee and Bogan, 1986) that these were un-
common shells in the Tennessee River system in aboriginal
times. Fifty valves of the Appalachian Monkey Face Q. sparsa
(Lea, 1841), a species generally associated with small tributary
streams of the upper Tennessee River drainage, occurred in
the archaeological sample. It is a rare species and remaining
populations appear limited to the unimpounded stretches of
the Powell and Clinch rivers in upper East Tennessee and
southwestern Virginia. Parmalee and Bogan (1986) reported
113 valves of Q. sparsa from Middle Woodland and Mississip-
pian components at the Clinch River Breeder Reactor Plant
site, Roane County, Tennessee and a single valve of this
species was recovered at the Starnes site, a historic Cherokee
farmstead along the lower Tellico River, Monroe County, Ten-
nessee (Parmalee and Klippel, 1984).

Cyclonaias tuberculata (Rafinesque, 1820): The Pur-
ple Warty-back is a widely distributed and locally common
mussel in Tennessee in both small and large rivers. As
evidenced by the quantity of valves recovered from aboriginal
sites, it was an abundant shell also in prehistoric times. For
example, Morrison (1942), in his analysis of shells from the
Pickwick Basin mounds (Tennessee River, northern Alabama),
commented that it ‘“..was extremely abundant in all the
mounds. It constituted one of the major fractions of the mussel
fauna that was used for food in building up the shell deposits.’
Although there appears to have been a viable population pre-
sent prehistorically in the West Prong Little Pigeon River, the
number of valves recovered at the McMahan site (74, <2%
of the total) suggests it was not abundant.

Elliptio Rafinesque, 1820: Shells of the Elephant’s Ear
Elliptio crassidens (Lamarck, 1819) and the Spike E. dilatata
(Rafinesque, 1820) have been recovered in considerable
numbers at aboriginal sites located along large rivers such
as the Tennessee (see Parmalee et a/., 1982). E. crassidens
is typically a large river species where it can become abun-
dant locally, but occasionally a few individuals will become
established in small- to medium-sized streams such as the
West Prong Little Pigeon River. The Spike, on the other hand,
is often the most abundant species present in small rivers.
Although there were three times the number of shells of E.
dilatata than E. crassidens in the McMahan site sample, sug-
gesting a predominance of small river habitat, combined they
accounted for <3% of the total.

Hemistena lata (Rafinesque, 1820): The Cracking Pear-

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS 169

ly Mussel was reported to have occurred in the Ohio,
Cumberland and Tennessee River systems. Ortmann (1918)
commented that ‘‘It is undoubtedly a rare shell;’’ in some
rivers such as the upper Clinch, however, it is locally com-
mon (Ahistedt, 1984). It appears to have been a rare species
in the West Prong Little Pigeon River during the time the
McMahan site was occupied as evidenced by the recovery
of only one valve.

Lexingtonia dolabelloides (Lea, 1840): The former
ecological environs of the Slab-side Mussel included shoal
areas of the Tennessee River downstream at least as far as
Pickwick Landing Basin in northern Alabama and its larger
tributaries in upper East Tennessee. Impoundment has
eliminated its habitat in the Tennessee River, and L.
dolabelloides is now limited to and is generally uncommon
in rivers such as the Duck, Clinch and Powell. Ortmann (1918)
observed that “...here we have a case where a swollen form
(dolabelloides) is found in the larger rivers, and a compressed
one (conradi) in the smaller stream, with the intergrades ex-
isting between them.” This condition was apparent in the
McMahan site material, where valves of this species com-
prised ca. 2.5% of the total, but the thick-shelled, swollen form
predominated.

Plethobasus Simpson, 1900: Combined, shells of
Plethobasus cooperianus (Lea, 1834), the Orange-footed
Pimple-back and P cyphyus (Rafinesque, 1820), the
Sheepsnose, totaled ca. 2.5% of the naiad sample. In Ten-
nessee the former species was considered an inhabitant of
the deep stretches of the Cumberland and Tennessee rivers
and their large tributaries. With reference to P cooperianus,
Ortmann (1918) stated that ‘‘l also found it in French Broad
River, at Boyd Creek, Sevier County, Tenn. Records from
‘Holston River’ probably refer to the Tennessee, at any rate,
it must be a rare shell above Knoxville’’ Only 11 valves of it
were identified while 46 specimens of P cyphyus, a shell that
can be found in small rivers as well as large, were recovered.
Valves of the Sheepsnose from the McMahan site appeared
intermediate between the typical large river form that is drawn
out posteriorly with a distinct row of pronounced knobs, and
the small river form with the radial row of knobs on the disk
poorly developed and nearly obliterated in some specimens.

Pleurobema Rafinesque, 1820: A total of 79 valves
(2.0% of the sample), representing four species in this genus,
were recovered in the sample. Three of these, P cordatum
(Rafinesque, 1820), Ohio River Pigtoe; P plenum (Lea, 1840),
Rough Pigtoe; and P rubrum (Rafinesque, 1820), Pyramid
Pigtoe, are generally considered large river, deep water
species that only rarely become established in small- to
medium-size streams. Of the approximately 40,500 valves (ca.
50 species) identified from 15 aboriginal sites in the
Chickamauga Reservoir (Tennessee River), those of these
three species of Pleurobema accounted for nearly 13% of the
total (Parmalee et a/., 1982). Although these and certain other
big river species are represented in the McMahan sample,
their limited numbers suggest the probability that stretches
of deep water habitat in the West Prong Little Pigeon River
were limited compared with greater riffle and shoal areas
typical of small- to medium-size rivers.

The fourth species of Pleurobema recorded from the
site, P oviforme (Conrad, 1834), the Tennessee Clubshell, is
restricted to the upper Cumberland and Tennessee River
drainages and is one that typically inhabits the smaller
streams and rivers. The taxonomic position of this species
is not entirely clear: it is characteristic of small rivers of the
upper Tennessee River drainage and probably represents P
clava (Lamarck, 1819) of the Ohio and lower Cumberland and
Tennessee rivers. Ortmann (1918) lists P oviforme argenteum
(Lea, 1841) as ‘‘...the compressed form of oviforme, peculiar
to the headwaters and other small streams. It also generally
attains a larger size than the typical oviforme, and is more
rhomboidal in outline. It is in Little Pigeon River, at Sevier-
ville, Sevier Co., TN., but not very well developed here, the
majority of the specimens belonging to oviforme:’ Ortmann
implied by this that the medium-sized river form P oviforme
closely resembled the upper Ohio River form of P. clava, but
he made note of the extreme shell variability, a condition ap-
parent in the McMahan site specimens.

Alasmidonta Say, 1818: Shells of two species represen-
tative of this genus were recovered at the McMahan site. One,
the Elk Toe Alasmidonta marginata (Say, 1819), is widespread
throughout the small streams and medium-size rivers of East
Tennessee. However, it appears to have been a rare shell
prehistorically in the West Prong Little Pigeon River as only
one right valve of a mature individual was recovered. The other
species, the Slipper Shell A. viridis (Rafinesque, 1820),
although not abundant (31 valves) suggests a former viable
population at this point in the river. Ortmann (1918) states that
it, A. (Pressodonta) minor Lea, 1845, is ‘‘A characteristic small
creek species, locally abundant. It is found all over the region,
but strictly avoids the medium-sized and larger rivers.’ He
recorded it from the Little Pigeon River at Sevierville.

Anodonta, cf. A. grandis (Say, 1829): Although at pre-
sent one of the most widely distributed and locally abundant
shells throughout impounded stretches of Tennessee rivers,
a slow current and mud/silt substratum most favorable to the
Common Floater was probably limited prehistorically. Of in-
terest is the statement by Ortmann (1918) that ‘‘No Anodonta
has ever been reported from the upper Tennessee region’;
however, he does make reference to two specimens (in the
collection of Bryant Walker) collected in a small pond near
the French Broad River eight miles above Knoxville. Bogan
(1980) identified a single valve of A. grandis, found as a burial
accouterment, from the Toqua site, Little Tennessee River,
Monroe County. In his treatment of the mollusks from Pickwick
Basin (Tennessee River), Morrison (1942) listed A. grandis,
along with four other species in the subfamily Anodontinae,
as ‘‘...present in small numbers only.’ No valves of A. grandis
were identified from the thousands of naiads recovered from
aboriginal sites along the Cumberland, Clinch and Tennessee
rivers in Middle and East Tennessee (Parmalee et a/., 1980,
1982; Parmalee and Bogan, 1986). Only one incomplete right
valve from the McMahan site suggests that A. grandis was
prehistorically a rare shell in the West Prong Little Pigeon
River.

Lasmigona Rafinesque, 1831: Lasmigona costata
(Rafinesque, 1820), the Fluted Shell, occurs in both large

170 AMER. MALAC. BULL. 6(2) (1988)

rivers like the Cumberland and Tennessee and in small- to
medium-sized rivers like the middle Duck and the upper
Powell and Clinch. Ahistedt (1984) noted that it ‘...is an ex-
tremely common species in the upper Clinch in Tennessee
and Virginia.’ Judging by certain extant unmodified stretches
of the West Prong Little Pigeon River (Fig. 2), assuming them
to be not unlike prehistoric conditions, it would seem this river
would have provided favorable habitat for L. costata. However,
only eight valves were recovered at the McMahan site. L.
holstonia (Lea, 1831), the Tennessee Heelsplitter, a species
often found locally abundant in small and/or headwater
streams, was also poorly represented at the site (5 valves,
3 individuals). All three were juveniles, the largest measur-
ing 35.5 mm total length. Ortmann (1918) recorded it for the
Little Pigeon River, Sevier County.

Actinonaias ligamentina (Lamarck, 1819): Prehistorically
the Mucket was widely distributed and common throughout
the major rivers in Tennessee such as the Clinch, Holston,
Tennessee, French Broad, and Cumberland. At present,
however, except for local populations in these rivers (primari-
ly the Holston), populations of A. /igmentina are restricted
mainly to the unimpounded upper stretches of the Clinch and
Powell rivers in East Tennessee. In archaeological context,
the percentage of shells of the Mucket varied from 7.5% of
those recovered in 15 sites in the Chickamauga Reservoir
(Tennessee River) (Parmalee et a/., 1982), and 13.5% at the
Clinch River Breeder Reactor Plant site (Parmalee and Bogan,
1986), to nearly 16% in two sites along the middle Cumberland
River (Parmalee et a/., 1980). The total of 148 valves,
representing 3.8% of all identified shells recovered at the
McMahan site, suggests a former viable population of this
mussel in the West Prong Little Pigeon River. A right valve
of a mature individual exhibited a high degree of polish on

its external surface; this modification possibly resulted from
its use aS some form of shaping or smoothing tool in the
manufacture of ceramic vessels.

Toxolasma lividus (Rafinesque, 1831): A total of 131
shells belonging to the genus Joxolasma were assigned to
the species T. lividus, the Little Purple. With respect to the
Toxolasma complex in this region, the comments of Ortmann
(1918) are appropriate: ‘‘What Lea has described as U.
moestus (from French Broad River, Tenn.) undoubtedly is this
[T. lividus]: | have specimens from Little Pigeon River (tributary
to French Broad), which are fully identical with moestus. U.
[Toxolasma] cylindrellus Lea (Duck River, TN.) is in shape ab-
solutely identical with T. lividium; however, it differs by paler
color of epidermis and nacre.” In light of these comments,
it is possible that some of the specimens from the McMahan
site are T. cylindrellus (Lea, 1868), assuming it is a good
species. Many valves of Toxolasma from the site still exhibited
a faded but uniform purple nacre. This small naiad appears
to have been fairly common prehistorically in the West Prong
Little Pigeon River.

Epioblasma Rafinesque, 1831: Seven species belong-
ing to this genus were represented in the molluscan sample
from the McMahan site, but combined the number of shells
totaled only 106, 3.0% of all identified valves. Three of these
species, Epioblasma arcaeformis (Lea, 1831), the Sugar
Spoon; E. haysiana (Lea, 1833), the Acornshell; and E.
stewardsoni (Lea, 1852), the Cumberland Leafshell, are now
considered extinct (Stansbery, 1971). The Yellow Blossom E.
florentina (Lea, 1857), represented at the McMahan site by
a single right valve of a male and identified as probably E.
f. form florentina based on the descriptions of Ortmann (1918)
and Bogan and Parmalee (1983), is probably close to extinc-
tion. The large river, nodular form of the Tubercled Blossom

Fig. 2. View of West Prong Little Pigeon River, north edge of Pigeon Forge, TN. Unmodified stretch of river, but at present poor mussel habitat.

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS 171

E. torulosa torulosa (Rafinesque, 1820), can also be con-
sidered extinct. Ortmann (1918) commented that E. arcae-
formis was found in large and medium-sized rivers and that
it was present in the French Broad River at Boyd Creek, Sevier
County. E. stewardsoni also occurred in shoal areas of the
larger rivers, but, unlike the once abundant E. t. torulosa, it
was apparently ‘‘A rare species’ (Ortmann, 1918).

Of the seven species of Epioblasma identified from the
site, valves of Epioblasma capsaeformis (Lea, 1834), the
Oyster Mussel, were the most numerous (42). This mussel
is at present locally abundant in the upper unimpounded
stretches of the Clinch and Powell rivers; it also can be found
in limited numbers in other small- to medium-sized rivers in
Middle and East Tennessee. Ortmann (1918) reported it from
the Little Pigeon River. Surprisingly, only one valve of the
Cumberlandian Combshell E. brevidens (Lea, 1831), was
recovered at the McMahan site; it was widely distributed and
locally common in medium-sized rivers such as the Big South
Fork Cumberland, Clinch and Powell in the Cumberland and
Tennessee River drainages of East Tennessee.

Lampsilis Rafinesque, 1820: A total of 385 valves of the
Wavy-rayed Lampmussel Lampsilis fasciola (Rafinesque,
1820), representing ca. 10% of the naiad sample, was
recovered at the McMahan site. Ortmann’s (1918) comment
that this species of Lampsilis is ‘practically everywhere in the
larger rivers as well as in smaller streams, but apparently more
abundant toward the headwaters’ is appropriate relative to
the West Prong Little Pigeon River. On the basis of the ar-
chaeological record, it was a very common shell at the time
the McMahan site was occupied. However, extensive naiad
samples from large rivers in East Tennessee indicate that L.
fasciola was rare, at least in the stretches near the sites:
Cumberland River, 2 sites, 7 specimens in a sample of 827
valves (.12%) (Parmalee et a/., 1980); Tennessee River, 15
sites, 3 specimens in a sample of 27,875 valves (.01%)
(Parmalee et a/., 1982); Clinch River, 1 site, 21 specimens in
a sample of 23,905 valves (.09%) (Parmalee and Bogan, 1986).

Seventy-nine valves of Lampsilis ovata (Say, 1817), the
Pocketbook, about 2% of the sample, were found at the
McMahan site. The ‘‘typical’’ shell of L. ovata is character-
ized by the distinct and sharp posterior ridge and, according
to Ortmann (1918), it is restricted to the larger rivers. However,
he points out (Ortmann, 1918) that ‘‘All along its range, and
chiefly above Knoxville, it is accompanied by the var. ventri-
cosa, and intergrades with it;’’ specimens examined from the
Little Pigeon River, Sevierville were identified by Ortmann as
L. ovata ventricosa. However, all valves from the McMahan
site complete enough to ascertain the angle of the posterior
ridge were L. ovata and not L. o. ventricosa (more rounded,
lacking the sharp-angled posterior ridge). Although less abun-
dant than L. fasciola, there appears to have been a viable
population of L. ovata in the West Prong Little Pigeon River
during aboriginal occupation of the McMahan site.

Lemiox rimosus (Rafinesque, 1831): A species of the
upper Tennessee River drainage, the Birdwing Pearlymussel
formerly inhabited shoals of the large rivers as well as small
streams, but it is at present restricted to local populations in
medium-sized rivers such as the Duck and upper Clinch and

Powell. Parmalee and Bogan (1986) recorded 623 valves
(2.6% of the total) of this small mussel in a sample of 23,905
shells from the Clinch River Breeder Reactor Plant site, but
only 24 (.09% of a total of 27,875 valves) were recovered from
15 sites reported from the Chickamauga Reservoir (Tennessee
River) by Parmalee et a/. (1982). In his study of mollusks from
the Pickwick Basin, Morrison (1942) reported L. rimosus
’’...throughout the mounds, but...nowhere in great abun-
dance.’ Ortmann (1918) considered it a rare shell and, except
for one local population in the Duck River (Maury County),
Ahlstedt (1984) also noted that it could not be found in any
great numbers. Prehistorically it must have been a rare
species in the West Prong Little Pigeon River as only eight
specimens were recovered in the McMahan site sample.

Ligumia recta (Lamarck, 1819): the Black Sandshell is
widely distributed from Pennsylvania to Minnesota south to
Oklahoma and Alabama (Burch, 1975); it inhabits primarily
medium-sized to large rivers where it may become locally
numerous. With the recovery of only two valves at the
McMahan site, it must have been a rare shell in the West
Prong Little Pigeon River during the time the site was oc-
cupied. The assumption can be made that in the case of the
Black Sandshell, like other species represented by only one
or a very few valves, individuals became established from time
to time but, for whatever reason(s), the river proved unsuitable
for the development of viable populations.

Medionidus conradicus (Lea, 1834): The Cumberland
Moccasin is a species endemic to the Upper Cumberland and
Tennessee River systems, and its distribution was character-
ized by Ortmann (1918) as ‘‘Very abundant in the headwaters
and in small streams generally, but quite rare in the larger
rivers.’ In a sample of 761 identified mussel shells from Cheek
Bend Cave, a multicomponent (Archaic-Woodland: ca.
7,000-1,000 BP) rockshelter site along the Duck River, Maury
County, valves of M. conradicus (100) comprised 13.1% of the
sample (Parmalee and Klippel, 1986). A total of 172 shells of
this species (4.5% of the sample) were recovered at the
McMahan site.

A study of species composition and abundance of ex-
tant naiad taxa in the West Prong Little Pigeon River adja-
cent to the McMahan site covered a two year period from June
1985 through May 1987. Results of this investigation will be
considered in more detail in this paper under PRESENT
NAIAD POPULATIONS: LITTLE PIGEON RIVER SYSTEM,
but in the case of aboriginal vs extant Medionidus specimens,
a brief comment here is appropriate. Only 12 individuals of
the Cumberland Moccasin were obtained (at muskrat feeding
stations) during this two year period. The right valve of each
was measured (mm): Range, 46.0 - 62.0; Mean, 55.23. Dur-
ing the initial identification process, it was noted that the en-
tire series of Medionidus from the site was made up of small
specimens. Length of the complete valves (N=29) was
measured (mm): Range, 24.5 - 42.5; Mean, 32.08. It appears
that individuals in the modern population of M. conradicus
reach a considerably larger mean size (55.23 mm, modern,
vs. 32.08 mm, archaeological) than did those from prehistoric
context; the largest specimen from the McMahan site had not
attained the size of the smallest individual recovered in

172 AMER. MALAC. BULL. 6(2) (1988)

1985-87. It is reasonable to assume the Indian would have
gathered the large individuals as well as the small had they
been present, so for whatever reason(s) the prehistoric popula-
tion of M. conradicus in the West Prong Little Pigeon River
consisted of individuals that did not attain the size of those
found in living populations.

Obovaria subrotunda (Rafinesque, 1820): Once
widespread throughout the Ohio, Tennessee and Cumberland
river systems, the range and population densities of the Round
Hickory Nut are now greatly reduced. This species is adapt-
able to both large river and small stream habitats. Ortmann
(1918) considered it rare in the upper Tennessee region, in-
cluding the small stream form O. subrotunda levigata (Raf-
inesque, 1820), in tributaries of the Tennessee, Holston and
French Broad rivers above Knoxville. It appears to have been
a rare shell in the West Prong Little Pigeon River as only nine
valves were recovered at the McMahan site.

Potamilus alatus (Say, 1817): The Pink Heelsplitter oc-
curs throughout the Mississippi drainage from Pennsylvania
south to Arkansas, Tennessee and Alabama (Burch, 1975).
Often an abundant shell locally in large and medium-sized
rivers, it occurs less commonly in small streams. Like the
preceding species, P alatus was an uncommon to rare mussel
(nine individuals) prehistorically in the West Prong Little
Pigeon River.

Villosa Frierson, 1927: A total of 852 valves, represent-
ing at least three species within this genus, amounted to
22.0% of all freshwater mussel shells identified from the
McMahan site. All are typical of small- to medium-sized
streams and locally they can occur in large numbers. For ex-
ample, Ahlstedt (1981) noted that Villosa perpurpurea (Lea,
1861) [probably a purple-nacre form or variety of V. trabilis
(Conrad, 1834)] was ‘‘common’’ in Copper Creek, VA.
Parmalee and Klippel (1984) found V. iris (Lea, 1829), the
Rainbow, and V. vanuxemensis (Lea, 1838), the Mountain
Creekshell, to be the two most common mussels inhabiting
the Tellico River, Monroe County, TN. Of the 1,125 specimens
recorded from this river, these two species of Villosa com-
prised 68.4% of the sample.

Villosa trabilis, the Cumberland Bean, is a small- to
medium-sized river species that is known from the upper Ten-
nessee and Cumberland River drainages, although its
distribution appears spotty. For example, it is one of the few
species still surviving as a viable population in the Obed River,
Cumberland County, TN on the Cumberland Plateau. Both
the Rainbow and the Mountain Creekshell are common and
widely distributed in the streams of East and, to a somewhat
lesser extent, Middle Tennessee; the latter species is one of
the few naiads that often becomes abundant in the head-
waters. Judging by the number of identifiable valves (See Table
2) recovered, V. vanuxemensis was the most common species
of Villosa in the West Prong Little Pigeon River during the
period of site occupation. However, all three taxa had well
established viable populations and their abundance in the ar-
chaeological record indicates former extensive stretches of
fast current and riffles with a substrate composed of cobbles,
gravel and coarse sand.

Cyprogenia stegaria (Rafinesque, 1820): The Fanshell

was once found rather sparingly throughout the upper Ten-
nessee and Cumberland rivers of Tennessee. It has been
poorly represented in some aboriginal molluscan faunas in-
cluding two recorded by Parmalee et a/. (1980) from the
Cumberland River and at 15 sites along the Tennessee
(Chickamauga Reservoir, Parmalee et a/., 1982). However,
Morrison (1942) reported that it was ‘‘..found in moderate
abundance, in nearly all the samples studied [from the
Pickwick Basin mounds. Tennessee River]. Ahlstedt (1984)
found C. irrorata to be a relatively common shell in the upper
Clinch River in Tennessee and Virginia. Ortmann (1918) noted
that ““..in the lower Clinch it is quite abundant’; at the Clinch
River Breeder Reactor Plant site valves of the Fanshell totaled
2,463, 10.3% of the total naiad sample (Parmalee and Bogan,
1986). Although the West Prong Little Pigeon River would ap-
pear to have been suitable for the establishment of a viable
population of this mussel, judging by the archaeological
species assemblage recovered and local populations that
presently exist in rivers such as the upper Clinch, the occur-
rence of only five valves of C. irrorata in the McMahan site
molluscan sample attest to its former rarity there.

Dromus dromas (Lea, 1834): Prehistorically the
Dromedary Pearlymussel was one of the most abundant shells
inhabiting the Cumberland and Tennessee River systems. Ap-
proximately 9,800 valves, comprising 35.25% of the naiad
sample from 15 sites in the Chickamauga Reservoir (Parmalee
et al., 1982), and 111 valves (13.42% of the sample) from two
sites along the middle Cumberland River in Tennessee
(Parmalee et a/., 1980) are two examples attesting to its former
abundance. Moreover, Morrison (1942), with reference to the
Pickwick Basin mounds, Tennessee River, northern Alabama
commented that ‘...dromas must have been very abundant
here previously. These specimens are of good size for the
species, and made up a major part of the total mussel fauna
gathered for food.’ Although not common at the McMahan
site (32 identified valves, <1.0% of the sample), apparently
a few individuals and possibly small populations became
established from time to time. Except for six shells of juveniles,
all specimens of D. dromas from the site were the typical big
river form, swollen with a large knob or lump on each valve.

Ptychobranchus Simpson, 1900: Shells of two species
belonging to this genus, Ptychobranchus fasciolaris (Rafin-
esque, 1820), the Kidneyshell, and P subtentum (Say, 1825),
the Fluted Kidneyshell, were recovered at the McMahan site
and together totaled about 16% of the naiad sample. However,
shells of the latter species made up slightly over 13%. Ort-
mann (1918) stated that the Kidneyshell was ‘‘widely and
uniformly distributed over the upper Tennessee region, but
nowhere in great numbers.’ After nearly 70 years this state-
ment is still a fairly accurate evaluation of its status in Ten-
nessee, although impoundment and increased pollution and
silting problems have brought about some changes. Recovery
of 124 shells of P fasciolaris, both juveniles and adults, sug-
gests a former viable population of this species in the West
Prong Little Pigeon River.

The most numerous shell recovered in the McMahan
site naiad sample was the Fluted Kidneyshell. A total of 508
valves were identified as Ptychobranchus subtentum; in ad-

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS 173

dition, of the nearly 1,000 indeterminate fragmented valves,
close to 200 of these could also have been referable to this
species judging by incomplete tooth/hinge line and fluted
posterior slope sections. P subtentum is an inhabitant of small-
to medium-sized streams of the upper Cumberland and Ten-
nessee River systems, becoming most abundant toward the
headwaters. It is, for example, a very common shell locally
in the unimpounded stretches of the Powell and Clinch rivers
in northeastern Tennessee and southwestern Virginia. At the
time the McMahan site was occupied, the Fluted Kidneyshell
was an abundant mussel in the naiad assemblage and, in
addition to its value as a food resource, the Indian utilized
(almost exclusively) shells of this species as some type of tool
(Fig. 3). Approximately 175 valves exhibited modification to

the posterior ventral margin; the shells appeared to have been
used as some form of scraper, the ventral edge of each hav-
ing been ground or worn down at an angle toward the posterior
end. Riggs (1987) illustrates two valves of Actinonaias ligamen-
tina, recovered at an early 19th century Cherokee farmstead
(Bell Rattle Cabin site, Monroe County, TN), that were modified
in a like fashion as those from the McMahan site. He attributed
the modified edges to the shells use as a potter’s tool; i.e.
the valves were used to scrape and smooth clay vessels
before they were fired. Harrington (1922) mentions that ‘‘..the
cherokee formerly used mussel-shells and a marine shell, pro-
bably some species of Cardium, for this purpose’ (pottery
smoothing tool). Shells of P subtentum from the McMahan
site were obviously preferred for this function as only three

Fig. 3. Modified shells from the McMahan site. Valve section (length, 27.0 mm) with two perforations (A); thin shell disc (diameter, 19.5 mm)
with center drilled and partially serrated edge (B); marine shell gorget (diameter, 34.0 mm), rattlesnake design (C); shell scrapers, Ptycho-

branchus subtentum (D, D,) and Ptychobranchus fasciolaris (E).

174 AMER. MALAC. BULL. 6(2) (1988)

Fig. 4. Widened and relocated channel of West Prong Little Pigeon River, Sevierville, TN, May 1967, looking upstream from U.S. 411 and 441
Highway bridge. McMahan site on left bank beyond bend in the river. Photo courtesy Tennessee Valley Authority.

valves of other species, one specimen of Elliptio dilatata and
two of P fasciolaris, were encountered that exhibited the
ground ventral margin.

PRESENT NAIAD POPULATIONS:
THE LITTLE PIGEON RIVER SYSTEM

The Little Pigeon River system flows generally north-
west from the Great Smoky Mountains National Park to its
confluence with the French Broad River (River Mile 27.4; 43.8
km: Fig. 1), ca. 8.0 km below Douglas Dam. The entire water-
shed, consisting of 914 km2, is in Sevier County, TN. Middle
Prong and Porters Creek join to form the Little Pigeon River;
downstream it is joined by Webb and Bird creeks, East Fork,
and Middle Creek and West Prong Little Pigeon River at
Sevierville (referred to as West Fork until ca. 1970). Principal
tributaries of the West Prong Little Pigeon River are LeConte
Creek, Roaring Fork, and Mill and Walden creeks. Total length
of the Little Pigeon River is 45.4 km, that of the West Prong
Little Pigeon River, 43.0 km. With minor exceptions the up-
per three-fourths of the drainage system flows in steep, nar-
row, mountain gorges, heading at elevations up to over
1,830 m at the southern boundary of the Great Smoky Moun-
tains National Park (Tennessee Valley Authority, 1964). With
the exception of the East Fork, tributaries of the Little Pigeon
and West Prong Little Pigeon rivers are now apparently devoid
of mussel populations. In light of the steep gradient, rapid cur-
rent, and bedrock and boulder substratum characteristic of
the majority of smaller streams making up this system, it is
doubtful whether viable and varied mussel assemblages ever

existed in all but the lower reaches of the Little Pigeon and
West Prong Little Pigeon rivers.

Although Sevier County, formed in 1794, is considered
predominantly rural, the past three decades have seen a
phenomenal growth in urbanization, especially as it relates
to the tourist industry. This has come about as the popularity
of the Great Smoky Mountains National Park continues to
escalate and the cities of Gatlinburg, Pigeon Forge and, to
a lesser extent, Sevierville enlarge and diversify their facilities
to accommodate the ever-increasing number of tourists. En-
vironmental degradation of the Little Pigeon River system also
continues to increase as a result of siltation, discharge from
waste water treatment plants, and trash in general. Surpris-
ingly, however, viable populations of several species of
endemic fishes, turtles and mollusks continue to survive in
very local areas in the lower stretches of the Little Pigeon
River, and particularly in the West Prong Little Pigeon River
in Sevierville. In the case of freshwater mussels, it seems even
more suprising that the greatest diversity of species (albeit
not large) and abundance in the Little Pigeon River system
can be found in a 1.0 km stretch of the West Prong Little
Pigeon River that was widened by the Tennessee Valley
Authority during the period from June 1967 to May 1968 (see
Figs. 4 and 5). Beginning at the TN Hwy 441 bridge (channel
width, 36 m), the width was expanded to 62 m at a point
152 m downstream for a distance of 1.9 km. In addition, the
mouth of the river was relocated ca. 0.6 km below its former
junction with the Little Pigeon River: this modification
eliminated two 180° bends and allowed discharge farther
downstream, thus eliminating extreme periodic flooding that

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS 175

Fig. 5. West Prong Little Pigeon River, looking downstream, during period of low water (July, 1986). McMahan site along right bank.

inundated the main business district and suburbs of
Sevierville.

During the period June 1985-May 1987, a total of 15
collecting trips were made in the Little Pigeon River in a
stretch from the TN Hwy 66 bridge in Sevierville to just below
the confluence with the West Prong Little Pigeon River, a
distance of ca. 0.9 km. A total of 118 specimens, represent-
ing 11 species, were recovered (Table 3); shells of Fusconaia
barnesiana, Lampsilis fasciola, Villosa vanuxemensis and V.
iris comprised 93.2% of the sample. The one individual of
Anodonta grandis (Say, 1829), the Common Floater, taken here
(shell length 85.5 mm) was the only specimen of this species
encountered during this study. Except for one individual and
a left valve of Elliptio dilatata (Rafinesque, 1820), the Spike,
found in the West Prong Little Pigeon River, one relic shell
(chalky, periostracum badly eroded) recovered in this stretch
of the Little Pigeon River was the only other example of this
species found in the river system.

Although several locations on the Little Pigeon River
from immediately below the confluence with the West Prong
Little Pigeon River to its mouth (confluence with the French
Broad River), a distance of ca. 7.5 km, were surveyed on six
occasions during this two year study, no freshwater mussels
were encountered. A substratum of shifting sand, private
homes and small businesses lining the east bank and
croplands and pastures adjacent to the west bank, plus the
last ca. 1.1 km above the mouth being impounded, probably
contribute the void in mussel populations. In his study of the
effect of rechanneling on the fish population of Middle Creek,
Sevierville, Etnier (1972) was of the opinion that substratum
instability and the decreased variability of the physical habitat
were the most significant factors responsible for changes in

Table 3. Species of freshwater mussels inhabiting the Little Pigeon
River, TN Hwy. 66 bridge to confluence with West Prong Little Pigeon
River, Sevier County, TN. Specimens obtained primarily from muskrat
feeding stations, June 1985-May 1987.

No. of % of
Species Specimens Specimens

Fusconaia barnesiana (Lea, 1838) 35 29.41
Pleurobema oviforme (Conrad, 1834) 3 2.52
Anodonta grandis (Say, 1829) 1 84
Lasmigona costata (Rafinesque, 1820) 1 84
Toxolasma lividus (Rafinesque, 1831) 2 1.68
Epioblasma capsaeformis (Lea, 1834) 1 84
Lampsilis fasciola (Rafinesque, 1820) 26 21.85
L. ovata (Say, 1817) 1 84
Villosa iris (Lea, 1830) 16 13.45
V. vanuxemensis (Lea, 1838) 33 27.73
Totals 119 100.00

the fish fauna. Widening and other modifications of the Little
Pigeon River in Sevierville by the TVA, plus the aforemen-
tioned conditions downstream, all contributed to reducing the
environmental quality of the river for most aquatic organisms.
Less than six specimens of Villosa iris and Lampsilis fasciola
were found in the Little Pigeon River at the Walnut Grove
Bridge in Sevierville (River Mile 6.7; 10.7 km); these were relic
specimens and the apparent paucity of naiads inhabiting this
stretch of the river could be due in part to urban development
along the banks at this point and upstream. No mussels were
found in the Little Pigeon River upstream from the southern
city limits of Sevierville, so with the possible exception of an
occasional individual becoming established, viable mussel

176 AMER. MALAC. BULL. 6(2) (1988)

populations in the Little Pigeon River are at present restricted
to the stretch between the TN Hwy 66 bridge and its con-
fluence with the West Prong Little Pigeon River. A small but
apparently stable population of V. vanuxemensis was found
inhabiting a ca. 0.2 km stretch of the East Fork, but this is
apparently the only naiad species living in this small tributary
stream.

As previously mentioned (see METHODS), emphasis
on surveying the molluscan fauna of the Little Pigeon River
system centered on that stretch of the West Prong Little
Pigeon River adjacent to the McMahan site. This was started
initially after noting a number of shells of endemic species,
along with large quantities of shells of Corbicula fluminea
(Muller, 1774), the Asiatic Clam, scattered along the bottom
and at muskrat feeding stations. It was felt that monthly
surveys for a period of time (as it turned out, two years) would
provide an accurate index to extant species and the relative
size of their populations still inhabiting the river, and a com-
parison of the present mussel assemblage with that from a
prehistoric context at the McMahan site.

No quantitative data were obtained for the species of
gastropods still inhabiting the Little Pigeon River system. Two
species, Leptoxis praerosa (most can be referred to the
smaller species/form, L. subglobosa) and Pleurocera parvum,
are locally distributed throughout the Little Pigeon River
system, including some of the smaller tributaries such as the
East Fork, but they appear most abundant in those stretches
of the Little Pigeon and West Prong Little Pigeon rivers sup-
porting viable mussel populations. /o fluvialis, P canaliculatum
and Lithasia verrucosa, taxa represented in the McMahan site
molluscan assemblage, have been extirpated from the Little
Pigeon River system. Campeloma sp. occurs in moderate
numbers in the silt/mud substratum in the West Prong Little
Pigeon River adjacent to the McMahan site, the only locale
where it has been noted. Two other species, Pseudosuccinea
columella (Say, 1817) and Physella gyrina (Say, 1821), have
been noted in some numbers under boards and other trash
caught in vegetation along the banks; these taxa could be
recent, or historic, additions to the molluscan fauna and their
numbers could well increase as they appear tolerant of low
water quality and a mud/silt substratum.

Table 4 provides a list of the naiad species and the
number of each collected in the West Prong Little Pigeon
River from June 1985 through May 1987. ‘‘Number of
Specimens’”’ reflects the quantity of paired valves collected
that were judged to be fresh or ‘‘recently dead” because they
either contained remains of soft parts or the shell had not yet
become heavily stained with algae, the periostracum was not
eroded (other than normal erosion of the beak), and the nacre
was not chalky. The only specimen of Alasmidonta viridis, the
Slipper Shell, encountered during the two year survey was
not included in Table 4 because, although paired, the valves
were badly eroded; this individual had probably been dead
‘or several years. The same was true of a right and left valve
(two individuals) of Cyclonaias tuberculata, the Purple Warty-
back; these valves were badly eroded and represent in-
dividuals that had died at least several years ago.

Shells of four species, Fusconaia barnesiana, Lamp-

Table 4. Species of freshwater mussels inhabiting the West Prong
Little Pigeon River, Sevier County, Tennessee. Specimens obtained
primarily from muskrat feeding stations, June 1985-May 1987.

No. of % of
Species Specimens Specimens

Fusconaia barnesiana (Lea, 1838) 689 45.39
Quadrula pustulosa (Lea, 1831) 2 13
Elliptio crassidens (Lamarck, 1819) 1 .06
E. dilatata (Rafinesque, 1820) 1 06
Pleurobema oviforme (Conrad, 1834) 132 8.69
Lasmigona costata (Rafinesque, 1820) 47 3.10
Toxolasma lividus (Rafinesque, 1831) 50 3.29
Epioblasma capsaeformis (Lea, 1834) 46 3.03
Lampsilis fasciola (Rafinesque, 1820) 330 21.74
L. ovata (Say, 1817) 53 3.49
Leptodea fragilis (Rafinesque, 1820) 1 06
Medionidus conradicus (Lea, 1834) 12 719
Potamilus alatus (Say, 1817) 21 1.38
Villosa iris (Lea, 1830) 103 6.78
V. vanuxemensis (Lea, 1838) 30 1.98

Totals 1,518 99.97

silis fasciola, Pleurobema oviforme and Villosa iris comprised
nearly 83.0% of all specimens recorded. Specimens of F.
barnesiana, a locally common species in numerous small
streams of the upper Tennessee River drainage, accounted
for 45.4% of the present naiad assemblage from the West
Prong Little Pigeon River. Prehistorically it appears to have
also been a common species in this stretch of the river; 347
valves identified from the McMahan site (9.2%) ranked it as
one of the four most numerous taxa in the assemblage. Of
the 13 species recorded from the Tellico River by Parmalee
and Klippel (1984), shells of F barnesiana amounted to 9.1%
of the total. P oviforme, another locally common shell in small-
to medium-sized rivers, totaled 8.8% and 8.7% respectively
in the Tellico and West Prong Little Pigeon river mussel
assemblages. Both V. iris and V. vanuxemensis exhibit viable
populations in the West Prong Little Pigeon and Little Pigeon
rivers, but the number of individuals from the West Prong ac-
counted for only 8.7% of the total number of specimens while
those from the Little Pigeon River amounted to 41.5%. V.
vanuxemensis is a species adaptable to medium-sized rivers
as well as small tributary and headwater streams, and one
that often becomes locally abundant; 48.2% of the mussels
(543 specimens) obtained by Parmalee and Klippel (1984)
from the Tellico River were this species.

One of the most numerous of the naiad species in-
habiting both the Little Pigeon and West Prong Little Pigeon
rivers is Lampsilis fasciola; individuals collected from both
rivers over the two year survey period accounted for approx-
imately 22.0% of all specimens in each. Nearly 10.0% of the
valves recovered from the McMahan site were those of this
species. At least five other taxa, Potamilus alatus, Lasmigona
costata, Lampsilis ovata, Epioblasma capsaeformis, and
Medionidus conradicus appear to be maintaining viable
populations in the West Prong Little Pigeon River, although
the latter species is rare. Of special interest is the occasional
establishment of an individual of a species generally

PARMALEE: LITTLE PIGEON RIVER MOLLUSCAN FAUNAS WAC

associated in the Mississippi or Interior Basin drainage: these
include Quadrula pustulosa (2 juvenile specimens, ca. 5 and
6 years of age, plus 2 relic right valves); Elliptio crassidens
(1 living adult, 2 relic pairs and 1 relic left valve); E. dilatata
(1 specimen, 1 left valve); Leptodea fragilis (1 specimen: shell
length 88.5 mm; left valve of a juvenile: shell length 43.4 mm).
Probably included in this category is Cyclonaias tuberculata,
based on the relic right and left valves previously mentioned.
Very possibly migratory host fishes, moving up the Little
Pigeon River from the French Broad River, provide the
mechanism for this dispersal. Thus far their numbers have
not become great enough to result in the establishment of
viable populations. Of the living taxa of freshwater mussels
reported here from the Little Pigeon River system, L. fragilis
is the only species that was not represented in the ar-
chaeological assemblage from the McMahan site.

SUMMARY

The prehistoric molluscan fauna of the West Prong Lit-
tle Pigeon River, Sevier County, Tennessee is one of the
richest and most diverse known for a small river in the upper
Tennessee River drainage. Archaeological salvage excava-
tions carried out periodically from June through December
1985 at the McMahan site, a late Mississippian (AD 1300-1600)
village and mound complex situated adjacent to the West
Prong Little Pigeon River, resulted in the recovery of ca. 7,400
identified aquatic gastropod shells (6 taxa) and 3,855
freshwater mussel valves (45 taxa). Shells of Leptoxis praerosa
and Pleurocera parvum composed 93% of the gastropod
specimens recovered. The naiad assemblage was dominated
by Fusconaia barnesiana, F. subrotunda, Lampsilis fasciola,
Villosa spp. and Ptychobranchus subtentum (ca. 65% of all
identified valves). Although several taxa represented in the
archaeological sample, e.g. F subrotunda, Elliptio crassidens,
Cyclonaias tuberculata, Pleurobema cordatum, and Dromus
dromas can inhabit the deep water of large rivers as well as
shallow small rivers (in some instances reflected by dif-
ferences in shell form), all species identified from the
McMahan site are known to occur in small- to medium-sized
rivers. However, approximately 30 of these reach their widest
distribution and greatest population densities in small- to
medium-sized rivers with normal depths of 1 m, a coarse
gravel/small cobble/sand substratum, riffles and swift
Current.

Ortmann (1925) concluded ‘‘..that originally there must

have existed a separation of two faunistic types in two different
drainage systems, a Cumberlandian River and an Interior
Basin River, and that subsequently these two systems became

connected, so that their faunas had a chance to mingle.’
He noted earlier (Ortmann, 1924) that ‘‘At the present time,

the distribution of the Cumberlandian Naiad fauna is markedly
discontinuous, being found in the upper Cumberland, the
upper Duck, and the Tennessee above the Mussel Shoals, but
not in the lower Cumberland, the lower Duck, and probably
also the lower Tennessee (downward from some point below
the Mussel Shoals, which has not yet been ascertained).”’ Of
those species whose origin has been determined with some

degree of certainty (e.g. Ortmann, 1925; van der Schalie,
1973), the naiad taxa represented at the McMahan site con-
sist of about 43% from the Interior Basin (Mississippian)
drainage and 57% from the Cumberlandian region (see Table
2). Former stretches of pool and riffle habitat in the West Prong
Little Pigeon River within close proximity of the McMahan site
apparently provided ideal conditions for the establishment of
an abundant and varied molluscan fauna. Naiad taxa whose
origin was the Interior Basin drainage reached the Little
Pigeon River system via the French Broad River.

Analyses of a sample of substratum taken from a
stretch of the West Prong Little Pigeon River that appeared
to provide the best mussel habitat, judging by the number
of live individuals and taxa observed during periods of low
summer water levels, was composed of the following parti-
cle sizes (after Wentworth, 1922): medium sand, 16.34%;
coarse sand, 66.87%; very coarse sand, 13.48%. The balance
was composed of small pebbles, granules, fine sand and very
fine sand. This type of substratum, whether in large uniform
expanses, e.g. 30 x 90 m2, or in small patches among large
cobbles or between layers of bedrock, provides the most
suitable habitat for present day molluscan populations. A river
habitat (Fig. 2) probably not unlike the present one adjacent
to the McMahan site, clear cut banks and channel widening
by TVA notwithstanding, existed in late prehistoric times and
supported a rich molluscan fauna that was heavily exploited
by the Indian.

Data on species distribution and population densities
of freshwater mussels inhabiting the Little Pigeon River
system were obtained from June 1985 through May 1987. The
primary source of quantitative data was obtained from shells
discarded by muskrats at feeding stations. Although the Little
Pigeon River and several tributaries that could have supported
mussel populations were surveyed, emphasis was placed on
aca. 1.0 km stretch of the West Prong Little Pigeon River ad-
jacent to the McMahan site. In spite of, or as a result of a
widening and straightening of the channel by TVA in
1966-1967, viable mussel populations of 11 species of mussels
still exist in this stretch in spite of continued severe degrada-
tion of the river environment. Occasionally individuals of other
naiad species (in this study, five taxa) become established
in the Little Pigeon and West Prong Little Pigeon rivers, but
apparently in such low numbers that viable populations are
unable to develop.

ACKNOWLEDGMENTS

A special note of appreciation is extended to Mr. James A.
Temple, Sevierville, former owner of the McMahan site property, for
permitting me to carry out archaeological salvage work during soil
removal operations. | am grateful to Mr. Richard R. Polhemus,
Research Associate, Frank H. McClung Museum, University of Ten-
nessee, Knoxville for the opportunity to analyze the faunal material
he recovered in the McMahan site mound and adjacent village area
and to incorporate the resulting data with those obtained by me for
this study. Dr. Jefferson Chapman, Curator of Archaeology, Frank H.
McClung Museum, was most helpful in analyzing the recovered
ceramics and providing archaeological background data, along with
Mr. Polhemus, on the McMahan site. On one or more occasions the

178 AMER. MALAC. BULL. 6(2) (1988)

following individuals assisted me with the archaeological field work
and/or collecting mollusks in the Little Pigeon River: Kenneth Can-
non, William Dickinson, Mary Ellen Fogarty, Jay Griswold, Richard
Kirk, Bruce Manzano and Lynn Snyder. | especially wish to thank
J. David Parmalee for his valued assistance in both the above
categories throughout the length of this study. Several other persons
contributed either pertinent data or special services and | wish to
thank them for their greatly appreciated assistance: Dr. Arthur E.
Bogan, Department of Malacology, Academy of Natural Sciences,
Philadelphia, PA (verification of several mollusks); J. Bennet Graham,
Division of Land and Economic Resources, Tennessee Valley Authori-
ty, Norris, TN (making accessable through TVA offices photographs
of the 1966-67 channel improvement project); Michael Morris, Depart-
ment of Anthropology, University of Tennessee, Knoxville, TN (analysis
of the West Prong Little Pigeon River substratum sample); Richard
E. Ruth, Tennessee Valley Authority, Knoxville, TN (supplying infor-
mation on Little Pigeon River channel improvement and other river
system data); W. Miles Wright, Frank H. McClung Museum (prepara-
tion of figures in this paper); Betty W. Creech, Frank H. McClung
Museum and Kim Johnson, Department of Anthropology, University
of Tennessee, Knoxville (typing drafts of the manuscript).

LITERATURE CITED

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Ahlstedt, S. A. 1984. Twentieth Century changes in the freshwater
mussel fauna of the Clinch River (Tennessee and Virginia).
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Bogan, A. E. 1980. A comparison of late prehistoric Dallas and Overhill
Cherokee subsistence strategies in the Little Tennessee River
valley. Doctoral Dissertation, Department of Anthropology,
University of Tennessee, Knoxville. 210 pp.

Bogan, A. E. and P. W. Parmalee. 1983. Tennessee’s Rare Wildlife,
Volume II: The Mollusks. Tennessee Wildlife Resources Agen-
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numbered). 321 pp.

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of a Portion of the Ethnologic and Archaeologic Collections
Made by the Bureau of Ethnology During the Year 1881. Third

Annual Report of the Bureau of Ethnology 1881-82, pp. 433-452.
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chaeological Survey of Pickwick Basin in the Adjacent Por-
tions of the States of Alabama, Mississippi and Tennessee.
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per Tennessee Drainage, with notes on synonymy and distribu-
tion. Proceedings of the American Philosophical Society
57:521-626.

Ortmann, A. E. 1924. The naiad-fauna of Duck River in Tennessee.
American Midland Naturalist 9(2):18-62.

Ortmann, A. E. 1925. The naiad-fauna of the Tennessee River system
below Walden Gorge. American Midland Naturalist 9(8):321-372.

Parmalee, P. W. and A. E. Bogan. 1986. Molluscan remains from
aboriginal middens at the Clinch River Breeder Reactor Plant
site, Roane County, Tennessee. American Malacological
Bulletin 4(1):25-37.

Parmalee, P. W. and A. E. Bogan. 1987. New prehistoric distribution
records of /o fluvialis in Tennessee with comments on form
variation. Malacology Data Net 2(1/2):42-54.

Parmalee, P. W. and W. E. Klippel. 1984. The naiad fauna of the Tellico
River, Monroe County, Tennessee. American Malacological
Bulletin 3(1):41-44.

Parmalee, P. W. and W. E. Klippel. 1986. A prehistoric aboriginal
freshwater mussel assemblage from the Duck River in mid-
dle Tennessee. Nautilus 100(4):134-140.

Parmalee, P. W., W. E. Klippel and A. E. Bogan. 1980. Notes on the
prehistoric and present status of the naiad fauna of the mid-
dle Cumberland River, Smith County, Tennessee. Nautilus
94(3):93-105.

Parmalee, P. W., W. E. Klippel and A. E. Bogan. 1982. Aboriginal and
modern freshwater mussel assemblages (Pelecypoda:
Unionidae) from the Chickamauga Reservoir, Tennessee.
Brimleyana 8:75-90.

Riggs, B. H. 1987. Socioeconomic variability in Federal Period Overhill
Cherokee archaeological assemblages. Master’s Thesis,
Department of Anthropology, University of Tennessee, Knox-
ville. 189 pp.

Stansbery, D. H. 1971. Rare and endangered mollusks in eastern
United States. In: Rare and Endangered Mollusks (Naiads) of
the U. S. S. E. Jorgensen and E. W. Sharp, eds. pp. 5-18.
Bureau of Sport Fisheries and Wildlife, United States Depart-
ment of the Interior, Twin Cities, Minnesota.

Tennessee Valley Authority. 1964. Sevierville, Tennessee Flood Relief
Channel Improvement Plan. Tennessee Valley Authority Divi-
sion of Water Control Planning, Flood Control Branch, Knox-
ville. Planning Report No. 0-6456. 46 pp.

van der Schalie, H. 1973. The mollusks of the Duck River drainage
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Date of manuscript acceptance: 4 September 1987.

EVALUATION OF TECHNIQUES FOR AGE DETERMINATION OF
FRESHWATER MUSSELS (UNIONIDAE)

RICHARD J. NEVES

AND

STEVEN N. MOYER’

VIRGINIA COOPERATIVE FISH AND WILDLIFE RESEARCH UNIT
DEPARTMENT OF FISHERIES AND WILDLIFE SCIENCES
VIRGINIA POLYTECHNIC INSTITUTE AND STATE UNIVERSITY
BLACKSBURG, VIRGINIA 24061, U. S. A.

ABSTRACT

Age validation and an assessment of four age determination techniques; shell ashing, thin-
sectioning, acetate peels, and enumeration of external growth bands, were conducted on several species
of freshwater mussels (Unionidae) in southwestern Virginia. The recovery of tagged and marked
specimens of four species after one to three years confirmed the formation of one distinct annulus
per year on and in shells. Thin-sectioning of valves was the most effective technique for aging and
provided a high degree of both accuracy and precision. Shell ashing was totally unreliable, and acetate
peels were inferior to thin-sections. The commonly used method of counting external growth bands
on shells consistently underestimated the ages of older specimens and is of limited use in age deter-

mination of unionids.

The determination of absolute ages of bivalves is
essential to derive population statistics for managing their
harvest and conservation. Shells (valves) of freshwater
mussels (Unionidae) exhibit pronounced bands or rings on
their external surface, and the distance between bands
decreases progressively with an increase in shell size. The
significance of these bands and their use to derive absolute
ages of mussels was discussed by early researchers (LeFevre
and Curtis, 1912; Isley, 1914; Coker et a/., 1921). Based on
the cyclical periodicity of band formation on valves, ages of
freshwater mussels have been determined using the tech-
niques of enumerating growth rings on the valve surface
(Chamberlain, 1931; Stansbery, 1961), and ashing shells in
a muffle furnace to separate the bands (Sterrett and Saville,
1975). The occurrence of growth bands within radial cross-
sections of the shell and hinge ligament has provided an ad-
ditional means of age determination (Hendelberg, 1960; Bjork,
1962; Ray, 1978; McCuaig and Green, 1983).

In most early attempts to age unionids, investigators
relied on the visibility of growth bands on the outer surface
of shells. Although these bands can be used to delimit age
of some species, in other species subjective and conflicting

1Present address: National Wildlife Federation, 1412 16th Street,
N.W., Washington, D.C. 20036, U.S.A.

ages typically result. Growth bands on lentic species, which
grow rapidly early in life, are characterized by regular spac-
ing and distinctness (Chamberlain, 1931; Stansbery, 1961),
whereas those on stream-dwelling mussels are less pro-
nounced (Grier, 1922; Brown et al., 1938). investigations to
determine age from external growth bands of riverine mussels,
hereafter called the growth ring method, is often hampered
by erosion of the shell surface, obscurity of bands on dark-
colored valves, subjectivity in distinguishing annuli from
stress-produced checks, and the inability to count closely
deposited bands near the valve margin of older specimens
(Ansell, 1968; Coon et a/., 1977; Lutz and Rhoads, 1980).
Population statistics derived from this method, which ap-
parently lacks both accuracy and precision, are therefore
fraught with problems.

In contrast to the growth ring method most often used
on freshwater bivalves, the techniques for determining ages
of marine bivalves have been rigorously tested and are ap-
parently more reliable. Most age studies of marine bivalves
since Barker (1964) have used two sectioning techniques, thin-
sections or acetate peels, to determine absolute ages; these
methods are now used routinely in marine malacology (Clark,
1980). Both the chondrophore and entire valve of marine
clams have proven to be useful for age determinations (Ropes
and O’Brien, 1980), and detailed descriptions of the methods

American Malacological Bulletin, Vol. 6(2) (1988):179-188

179

180 AMER. MALAC. BULL. 6(2) (1988)

are provided by Lutz and Rhoads (1980) and Ropes (1984).

The annual formation of winter growth bands on the
valve surface of some freshwater mussel species has been
documented (Isley, 1914; Chamberlain, 1931; Negus, 1966;
Haukioja and Hakala, 1978), but the formation of internal an-
nuli lacks appropriate verification. Most studies that have
estimated ages of mussels by these various methods typically
omit age validation (i.e. proof of the accuracy of the technique).
Validation of these methods for mussels is necessary because
of the presence of less prominent, stress-related growth
checks in bivalve shells, termed pseudoannuli or ‘‘false’’ an-
nuli. Some researchers have been able to distinguish the dif-
ference between annuli and ‘‘false’’ annuli with relative ease
(Chamberlain, 1931; Negus, 1966; Day, 1984); others have had
difficulty, especially with riverine species (Coon et al., 1977;
Haukioja and Hakala, 1978). Previous studies with unionids
in the upper Tennessee River drainage, of Virginia and Ten-
nessee, have also experienced difficulty in delimiting annuli
and recognized the need for validation (Zale, 1980; Weaver,
1981). Age validation is an essential prerequisite for obtain-
ing sound population statistics, and the application of routine
but unvalidated methods to all species can result in signifi-
cant misinterpretations of biological data (Beamish and
McFarlane, 1983a, 1983b).

The three objectives of our study were: (1) validation
of the annual formation of growth bands on and in the valves
of various sizes and species of unionid mussels; (2) tests of
the utility of shell ashing, thin-sectioning, and acetate peels
for freshwater mussels; (3) comparison of the ages of
specimens derived from the growth ring and thin-sectioning
methods.

MATERIALS AND METHODS

ANNULUS VARIATION

A mark and recovery program was conducted from
1979 to 1983 to validate the annual deposition of growth bands,
to determine the season of annulus formation, and to provide
empirical data on mussel growth. Four relatively common
mussel species, representing three subfamilies of unionids,
were selected for this phase of the study: Pleurobema oviforme
(Conrad, 1834); Lasmigona subviridis (Conrad, 1835); Villosa
vanuxemi (Lea, 1838); and Medionidus conradicus (Lea, 1834).
Specimens were obtained from three sites in western Virginia:
New River, Montgomery County; North Fork Holston River,
Smyth County; Big Moccasin Creek, Russell County. A total
of 1452 adult mussels were collected by hand, transported
to our laboratory, and held in a 300 / aerated, recirculating
tank (Table 1). Each specimen was measured (length and
height) with calipers to the nearest 0.1 mm and marked by
one of three methods, numbered tag only, tag plus valve
notch, and tag plus painted valve. These marking methods
were used to record shell growth for a known time period and
to recognize differences between annuli and other bands
(false annuli) formed externally and internally on the valves.

One valve of each mussel was tagged with a3 x 5mm
fluorescent orange, sequentially numbered disc tag (Floy Tag
Company, Seattle, Washington), held in place by Duro

superglue (Loctite Corporation, Cleveland, Ohio). A small
triangular notch was filed in the ventral margin of notched
specimens, and red fingernail polish was applied to the shell
margins of painted specimens. The marked specimens were
transplanted to two sites (| and II) in each stream; specimens
at site | (15 to 25 m2 in area) were tagged and 7% were
painted, and those at site II (0.7 to 3 m2) were tagged and
notched (Table 1). Mussels were returned to their collection
sites within 2 weeks and placed, properly oriented, in the
substratrum. At site | in the New River, 150 tagged mussels
were divided among three substrata-filled chicken wire
enclosures (18 mm mesh; 76 x 76 x 13 cm) set into the
substratrum to inhibit mussel dispersal and facilitate periodic
examination. The remaining mussels at this site were placed
near the enclosures. Sites in all three streams were identified
either by landmarks, streambed features, or markers.

In each stream, mussels at site | were recovered after
1 year for annulus validation, and a sample of about 12
mussels at site Il was collected quarterly during the first year
for examination of seasonality in growth band deposition.
Some specimens that could not be found 1 year after plant-
ing were collected up to 4 years later (1983). Recovered
mussels were sacrificed, and incremental growth on valves
was measured and examined for annulus formation externally
and internally, under a dissecting microscope.

EVALUATION OF AGE TECHNIQUES

Ashing of shells to separate growth layers followed pro-
cedures similar to those used by Sterrett and Saville (1974).
Initial cuts made on a Buehler Isomet low-speed saw unit with
a diamond-impregnated blade (Buehler Ltd., Evanston, Illinois)
were: (1) from the umbo to the shell margin along the vector
of maximum length, and (2) from the umbo to the shell margin
perpendicular to the first cut. The triangular wedges of shell
produced by these cuts, with sectioned surface exposed on
two sides, were ashed in a muffle furnace. Sterrett and Saville
(1974) recommended ashing at either 500°C for 10 minutes
or 600°C for 5 minutes. Because temperature and time are
the factors apparently crucial for producing good results, a
size range of shells (20-80 mm) was ashed at both of the
recommended times and temperatures. However, the resulting
ashed shells were too brittle to allow effective separation of
many of the growth layers. Therefore, we conducted a series
of ashing time and temperature trials to evaluate the utility
of this technique: 300°C for 1, 5, 10, 15, or 20 min; 400°C for
1, 5, 10, or 15 min; 500°C for 1, 5, or 10 min; and 600°C for
1 or 5 min. Preliminary ashing tests indicated that each of
these combinations of times and temperatures could produce
usable results. Three shells, small (<40 mm), medium (40-60
mm), and large (> 60 mm), were ashed in each of the 14 trials.
All trials were later replicated to corroborate initial results. Utili-
ty of the shell ashing technique was assessed by (1) how well
annual layers could be separated, and (2) how well growth
bands could be distinguished externally and in cross-section.

Thin-sectioning of valves followed procedures similar
to those described by Clark (1980), in which a low speed saw
unit and diamond-impregnated blade was used. An initial

NEVES AND MOYER: AGING OF FRESHWATER MUSSELS 181

Table 1. Number of mussels of four species tagged in 1979 - 1982 at two sites each on Big Moccasin
Creek (BMC), North Fork Holston River (NFHR), and New River (NR), western Virginia.

Stream, Site Pleurobema Medionidus Villosa Lasmigona
and Date oviforme conradicus vanuxemi subviridis Total
BMC |

Oct 1979 — 63 29 _ 92

Oct 1980 39 2 6 — 47

Sept 1981 101 165 103 _— 369
BNC II

Jul 1982 2 35 12 — 49
NFHR |

Sept 1981 152 139 108 — 399
NFHR II

Jul 1982 30 27 4 — 98
NR |

Apr 1982 — — — 320 320
NR Il

Jul 1982 — — _- 78 78
Total 324 431 299 398 1452

cross-sectional cut from the umbo to the shell margin followed
the vector of maximum growth (posterio-ventrally), since it
generally intersected growth lines at right angles. Shell cuts
were then bonded to petrographic micro-slides (27 x 46 mm)
with epoxy glue (Buehler epo-kwick) and vacuum-sealed in-
to a petrographic chuck attached to the cutting arm of the
saw. Because the thickness of the second cut was critical to
producing thin-sections of suitable quality, several cuts rang-
ing from 200 to 380 um were made to determine optimal
thickness for growth band detection. A thickness of 280 um
was considered to be best for consistent, high resolution thin-
sections and was used in all subsequent sectioning of valves.
The utility of thin-sectioning was evaluated on a varie-
ty of mussel species from rivers in southwestern Virginia. Shell
lengths ranged from 15 mm for Medionidus conradicus to 210
mm for Potamilus alatus (Say, 1817), although most shells were
20 to 80 mm long. Shells longer than 60 mm had to be cut
more than once because the saw blade was only 114 m in
diameter. The final cut through the umbonal region of large
shells included all internal growth lines. Sectioned shells and
derived thin-sections were examined under 4X magnifica-
tion, and felt-tip pen marks were made adjacent to the point
where each growth line exited at the shell surface. The cross-
sectioned shell was then superimposed on the marked thin-
sections. This justaposition of shells allowed for visual com-
parison of internal with external growth lines to corroborate
contiguity and to identify false annuli on the valves.
Acetate peels from sectioned shells followed the
method of Kennish et al. (1980). Shells of Pleurobema oviforme,
Medionidus_ conradicus, Villosa vanuxemi, as well as
Fusconaia cor (Conrad, 1834) and F. cuneolus (Lea, 1840),
two federally endangered species, were separated into small
(<40 mm), medium (40-60 mm), and large size groups (>60
mm). An initial cross-sectional cut was made with the low-
speed saw from the umbo to the shell margin along the vec-

tor of maximum growth. Although Kennish et a/. (1980) sug-
gested pre-embedding the valves in an epoxy resin first to
prevent fracturing during sectioning, the stability of the low-
speed saw allowed sectioning of most shells without fracture
(Clark, 1980). Valve sections were then ground by hand on
sequentially finer grit sizes: 320, 400, and 600 (Buehler car-
borundum grits) and polished with polishing alumina (Fisher
Scientific Co., Fairlawn, New Jersey) on felt polishing cloth.
Because acid-etching is the critical step in this technique and
is apparently related to shell structure, mineralogy, organic
content, and state of preservation (Kennish et al. , 1980), etch-
ing times and HCI concentrations are expected to differ slightly
among species. Therefore, polished sections of each species
and size group were etched in a dilute solution of HCI at
various concentrations (1%, 5%, 10%) and time periods (15
sec to 5 min). This allowed development of an optimal pro-
cedure for shells of a given size and species. One valve was
used in each of the etching time and HCI concentration trials.
The etched shell sections were washed under running water
and dried.

In the last step of the peel process, we placed the
etched section firmly on a strip of acetate (2 mm thick) covered
with acetone, and pressed for 30 sec. After the acetone dried
completely (2-3 hr), the valve was pulled from the acetate strip,
leaving an imprint (the peel) on the acetate. Internal growth
bands on the peel were counted under 4 to 10X magnifica-
tion. Quality of the acetate peels was judged by two criteria:
clarity of growth bands in the umbonal region, and degree
to which bands could be traced from the umbo to the shell
margin.

COMPARISON OF EXTERNAL AND

INTERNAL AGES

The valves of 82 specimens of Fusconaia cor and
Pleurobema oviforme were selected for this comparison.

182 AMER. MALAC. BULL. 6(2) (1988)

These species had relatively distinct external growth bands
and were aged by the growth ring method. Later, the same
valves were thin-sectioned, as previously described. Ages
determined by these two methods were plotted graphically,
and a Wilcoxon signed rank test was used to compare
differences.

RESULTS

ANNULUS VALIDATION

A total of 521 (36%) of the 1452 marked mussels was
recovered from the three streams (Table 2). Recovery rates
of specimens from Big Moccasin Creek and the North Fork
Holston River were similar, 49.1 and 47.1% respectively; the
largest species, Pleurobema oviforme, was the most frequently
recovered. Both sites on the New River yielded low returns
(3.2%) because of specific problems. Muskrats (Ondatra
zibethicus L. along the New River removed 55 marked spec-
imens (found in shell middens) in June-July 1982, and
one enclosure of 50 mussels was vandalized in October. In
addition, a thick mat of Elodea developed by fall 1982 and
summer 1983, and caused considerable siltation and mortality
of marked mussels.

Of the three marking methods tested, notching of

Fig. 1. Thin-section of the umbonal region of Pleurobema oviforme
showing internal growth lines (bar = 1 mm).

Table 2. Recovery and validation of annulus formation on mussels
marked in Big Moccasin Creek (BMC), North Fork Holston River
(NFHR), and New River (NR), western Virginia.

Stream/Species Mussels
Recovered No.
No. % validated
Big Moccasin Creek
Pleurobema oviforme 83 58 7
Medionidus conradicus 101 38 10
Villosa vanuxemi 90 60 9
Subtotal 274 49 26
North Fork Holston River
P. oviforme 109 60 4
M. conradicus 63 38 9
Vv. vanuxemi 62 42 20
Subtotal 234 47 33
New River
Lasmigona subviridis 13 3 4
Total 521 63

valves was the most useful for recording shell growth and an-
nulus deposition. Annuli appeared as dark bands in sectioned
valves (Clark, 1974; Lutz and Rhoads, 1980), and were evi-
dent on 25 (27%) of the 94 notched specimens recovered at
site Il in the streams. Notching readily identified the origin
of incremental growth and subsequent growth at the shell
margin (Fig. 1). Thin-sections through the notch clearly
delineated incremental growth and the presence of a growth
band. An annulus was validated on all notched shells that
grew more than 1 mm/yr and on several shells that grew 0.5
to 1.0 mm/yr. Several specimens, marked between 1979 and
1982 and collected in 1983, showed one annulus for each year
at large.

Although the disc tags remained firmly attached to all
specimens upon recovery, mussels with only tags were less
useful for documenting growth bands. Only 38 (9%) of 425
recovered specimens from site | in the streams were useful
for annulus validation. All mussels that grew more than 1.5
mm/yr were validated, but lack of precision with caliper
measurements and a fragile shell margin prevented annulus
validation on a higher percentage of the slower-growing
specimens. Fingernail polish on shell margins was completely
ineffective. Within 3 months after marking, it had sloughed
from the shells apparently due to abrasion in the substratum.

Annulus formation was documented on 63 (12%) of the
521 specimens recovered from all sites (Table 2). Although
this percentage appears low, only specimens with readily
measurable incremental growth in length (1.0-1.5 mm, de-
pending on marking method and species) could be used for
validation. Occurrence of single (annual) growth bands was
confirmed in the shells of all four marked species. Because
83% of the recovered specimens grew less than 1 mm, growth
bands formed during the last year on these mussels were
nearly indistinguishable from those formed during the
penultimate year (Table 3). Growth was most rapid in
Lasmigona subviridis, the most thin-shelled species, whereas

NEVES AND MOYER: AGING OF FRESHWATER MUSSELS

183

Table 3. Annual growth increments on mussels tagged and recovered in Big Moccasin Creek, North Fork Holston River, and New River, western

Virginia.
Stream and Species (0-<1) (1-< 2)
No. % No. %
Big Moccasin Creek
Pleurobema oviforme 67 81 12 15
Medionidus conradicus 92 91 6 6
Villosa vanuxemi 71 79 18 20
North Fork Holston River
P. oviforme 98 90 11 10
M. conradicus 61 97 2 3
V. vanuxemi 51 82 11 18
New River
Lasmigona subviridis 42 57 20 27
Total 482 83 80 13

adults of the other species grew more slowly.

Despite the slow growth of most of the recovered
mussels, validation results provided convincing evidence of
the formation of a single growth band each year. An annulus
was formed in all tagged specimens that grew more than
1.5 mm and all tagged and notched specimens that grew more
than 1.0 mm during the year. None of these lacked an an-
nulus, nor had they more than one prominent growth band.

Only limited evidence was obtained on the seasonali-
ty of annulus formation, primarily because growth was slow
throughout the year. Some specimens from the three streams
provided evidence that the growth band had formed between
January and May. No annulus was observed on specimens
examined during fall and winter, but valves of four mussels
examined in May and all of 16 valves examined in July had
an annulus within the outer layer of incremental growth.
Although sample sizes are small, it appears that the annulus
is formed (becomes visible) in spring in western Virginia.

EVALUATION OF AGE TECHNIQUES

All ashing trials failed to meet our two criteria for
Suitability in age determination; i.e. separation of each annulus
and recognition of growth bands externally and internally.
Shells were either too brittle or inseparable at many annuli
after the tests. Most shells ashed at 400°C for 10 and 15 min
did separate along the first one to four annual growth bands.
However, subsequent annuli could not be separated con-
sistently; shells were brittle and crumbled when manipulated.
Ashing also tended to obliterate the recognition of growth
bands, making true annuli and false annuli indistinguishable.

The acetate peel technique was less effective than thin-
sectioning, both in terms of clarity of growth bands in the um-
bo region and degree to which bands could be traced
throughout the shell. Because of the similarity of the thin-
section and peel techniques, and higher resolution produced
by thin-sectioning, acetate peels produced by the method
described were judged to be inferior to thin-sections for deter-
mining ages of mussel shells.

Annual increment (mm)

(2-<3) (3-< 4) (4-<5)
No. % No. % No. %
2 2 1 1 1 1

2 3 — — —

1 1 — — — —

9 13 2 2 1 1
14 3 3 <1 2 <i

Thin-sectioning of valves was the most effective tech-
nique and usually provided a high degree of precision (Fig. 1).
A section thickness of 280 um produced consistent, high quali-
ty preparations for valves of all species over a wide range of
shell lengths (15-210 mm). Ages of sectioned shells ranged
from 3 to 56 years. The clarity of thin-sections resulted in a
high degree of accuracy because the contrast between true
annuli and false annuli was pronounced, and annuli could
usually be traced continuously from the umbo to the shell
margin. The entire sectioning procedure required 0.5 to 1 hour
per valve (excluding overnight hardening of the epoxy glue),
depending on shell size and thickness.

RECOGNITION OF ANNULI

Species that displayed distinct external annuli also had
distinct internal annuli. Shells of Pleurobema oviforme and
Fusconaia cor typically had well-defined internal and exter-
nal annuli, unlike those of F cuneolus, Medionidus conradicus
and Lasmigona subviridis. Internal annuli of Villosa vanuxemi
were readily distinguished, but the external growth bands were
obscured by the dark periostracum of this species. Signifi-
cant variability in the clarity of external annuli was also evi-
dent within a species; erosion of the shell surface was the
major contributing factor, and this problem was directly cor-
related with age. Young specimens (3-6 yrs) were rarely af-
fected, but in older individuals (7-15 yrs), the first and often
second annulus was eroded. The first two annuli were typically
missing in the oldest specimens (>15 yrs), and those older
than 20 years could not be aged externally because the
periostracum had become extensively damaged. Shell cor-
rosion (dissolution) was also evident on shells from all three
streams. Prior dissolution of calcium carbonate in the umbonal
region apparently resulted in pit formation.

False annuli occurred occasionally in all species ex-
amined. Thin-sectioning provided the best method for identi-
fying false annuli because true annuli could be traced from
umbo to shell margin. In contrast, false annuli were
characterized by an incomplete growth line in thin sections
(Fig. 2). Recognition of false annuli was much more difficult

184 AMER. MALAC. BULL. 6(2) (1988)

Fig. 2. Thin-section of a valve of Pleurobema oviforme with a false
annulus (arrow) among true annuli (bar = 0.5 mm).

on the shell surface. For example, the inclusion of small par-
ticles from the substratum into shells often caused the for-
mation of a false annulus. This false growth check was ob-
served most commonly in shells of females, particularly in
Villosa vanuxemi from Big Moccasin Creek and the North Fork
Holston River. Incorporation of these particles in the shell pro-
duced a thick, dark line both internally and externally on the
shell (Fig. 3). This growth check appeared to be a true an-
nulus on the shell surface, but was not continuous in the cross-
sectioned shell.

EXTERNAL VERSUS INTERNAL AGES

Growth bands on the external surface of valves of
Pleurobema oviforme and Fusconaia cor were readily visible
and were more distinct than those in most other species
available for such a comparison. Annuli were easily discerned
on specimens 3 to 8 years old, but became more tightly
grouped and less distinct on valves of mussels 8 to 15 years
old. Shells of mussels more than 15 years old were difficult

Fig. 3. Thin-section of a valve of Villosa vanuxemi with the incorpora-
tion of sediment (arrow) into the valve (bar = 1 mm).

to age because surface annuli were nearly contiguous or in-
distinguishable even under magnification. If the periostracum
was damaged by erosion or corrosion on older specimens,
frequently no age estimates were possible. Erosion of valves
was especially prevalent on old specimens of P oviforme. No
valves older than 20 years, as determined by the thin-section
method, could be aged by the growth ring method because
of periostracum damage. Erosion was also the probable cause
for loss of the first and often second annulus on some valves
older than age 6 years. The thin, organic-rich growth checks
apparently were less solid than the calcium carbonate deposi-
tion in annual growth, and shell fractures in young specimens
were occasionally evident along the annulus. However,
cleavage lines were nearly always visible on the shell and
were counted as annuli.

A comparison of ages derived by counts of external
annuli and by thin sectioning on 82 specimens of Fusconaia
cor and 49 Pleurobema oviforme indicated that counts of ex-
ternal annuli consistently yielded underestimates of ages (Fig.
4). Differences in ages determined by the two methods were
highly significant (P<0.01). The degree of underestimation
was directly proportional to age estimates; the older the
specimen, the greater the underestimate of age by the growth
ring method. The two methods yielded similar ages for F.
cor up to age 10, but mussels 11 to 25 years old were under-
estimated by 1 to 5 years when external annuli were counted.
Thin-sectioning was more effective, particularly on old
specimens (> 20 yr). Eight valves of P oviforme older than
20 years could not be aged externally due to periostracum
damage; these specimens ranged in age from 25 to 56 years
based on thin-sections.

On thin-sections of the latter two species, marks were
made adjacent to the exit location of each annulus at the shell
margin to allow visual comparisons with cross-sectioned
shells from which the thin-sections were cut. Comparison of
the two clearly corroborated the occurrence of one external-
ly visible annulus with its internal counterpart in every shell.
This external-internal comparison also demonstrated the oc-
casional presence of thinner, false annuli on the shell sur-
face that had no counterpart internally. Generally, internal an-
nuli were much easier to distinguish than external annuli,
especially near the shell margin of older specimens.

DISCUSSION

Deposition of one prominent growth band annually was
validated in 12% of the tagged specimens that were recovered
from the three study streams. The relatively low recovery rate
(36%) and slow growth (<1 mm) of most specimens limited
the availability of a larger sample size. Negus (1966) recovered
only 56 (9.7%) of 572 marked specimens of three freshwater
mussel species in the Thames River, England after 1 year
to validate annulus formation; of these, 43 (77%) showed an
annulus. Although recovery rates of marked bivalves have
been typically low in both freshwater and marine environments
(Murawski et a/., 1982; Schaul and Goodwin, 1982), forma-
tion of annual growth bands in bivalves from temperate
climates appears to be common. In the tropics, unionids also

NEVES AND MOYER: AGING OF FRESHWATER MUSSELS 185

20 Fusconaia cor

15

10

20 Pleurobema oviforme

N = 49

GROWTH RING AGE (yr)

15

10

oO 5 10 15 20 25
THIN-SECTION AGE (yr)

Fig. 4. A comparison of age estimates for two species aged by the
thin-sectioning and external growth ring methods. Data points below
the 45° line represent underestimates of specimen ages by the growth
ring method.

exhibit shell bands, but the causes for their formation are pro-
bably different from those for temperate species (McMichael,
1952). This apparent regularity in banding could lead some
investigators to assume that annulus formation is a universal
phenomenon and that age validation might not be necessary.
However, we caution that annual periodicity of growth line
deposition is a hypothesis that should be confirmed for each
species and locality before it is accepted.

The slow growth of most tagged specimens (96% grew
less than 2 mm per year) was the major handicap in age
validation. Growth increments along the shell margin of these
specimens were insufficient to allow clear separation of growth
during the year after tagging from growth in the penultimate
year. Ages of most of the tagged specimens, determined later
by thin-sections, were 8 to 20 years. These older, larger

specimens proved to be unsuitable, in retrospect, for this com-
ponent of the study. Our age validation efforts were most suc-
cessful with mussels of the relatively faster growing, younger
age-classes. Therefore, a range of size classes of sufficient
number should be used in age validation to overcome the dif-
ficulties posed by the slow growth of adults of riverine species.

Other problems associated with slow growth included
accuracy of caliper measurements and growth layer detach-
ment. Unnotched mussels that grew less than 1 mm per year
had to be excluded because rough shell margins contributed
to measurement error with calipers, and annulus deposition
could not be confidently ascertained. The narrow growth band
along the shell margin often became brittle after the
specimens were killed and occasionally broke during
measurement or thin-sectioning. Despite these problems with
age validation, successes and failures provided experience
that improved precision in age determinations of shells. For
shells that grew sufficiently for measurement during the 1 year
period, the formation of a single growth band per year was
confirmed. The identification of both internal and external
growth bands for a specimen facilitated the recognition of true
versus false annuli and contributed to our confidence in age
determinations.

As judged by counts of annuli on mussel shells and
growth measured for up to 4 years at study sites, adults of
riverine species in Virginia grow slowly and reach maximum
ages greater than those reported for lentic species (Grier,
1938; Stansbery, 1961). Longevities of the species aged by
thin-sections ranged from 22 to 56 years. These ages exceed
those reported for some species in the Mississippi River (Coon
et al., 1977), are less than the extreme age (> 100 yr) reported
for Margaritifera margaritifera L. in Europe (Hendelberg, 1960),
but are apparently similar to ages of other slow-grewing
species (Isley, 1914; Stansbery, 1971). Isley (1914) and Coker
et al. (1921) reported that light-shelled species grow rapidly,
and subsequent studies on Anodonta spp. and other thin-
shelled species have confirmed their observations (Stansbery,
1961; Negus, 1966; Haukioja and Hakala, 1978). In com-
parison, they noted that growth in length of heavy-shelled
species is most rapid in early life but slows considerably, such
that growth lines become tightly spaced and difficult to dif-
ferentiate. Coker et a/. (1921) computed mean growth rates
of roughly 6 mm/yr for medium-sized individuals of thick-
shelled species (Quadrula spp.), and Isley (1914) observed
shell growth to be roughly 1 mm/yr for older (larger), riverine
individuals. Riverine populations of at least some mussel
species therefore contain many older, slow-growing cohorts.
Based on the slow growth, closely spaced annuli, and con-
siderable longevity of mussels, it is imperative that specimens
be accurately aged if exploitation potential or population
Statistics are to be assessed from age-class structure and
abundance (Moyer, 1984).

Although the formation of growth bands is the key pro-
cess that allows age determination, it is not well understood.
Band patterns on freshwater mussel shells occur in two
varieties, wide, dark bands at fairly regular intervals, and
lighter bands that are irregularly spaced (Tevesz and Carter,
1980). The mechanism through which these bands are incor-

186 AMER. MALAC. BULL. 6(2) (1988)

porated into the mussel shell is still unclear. Explanations for
this mechanism have been put forth by several authors, and
were reviewed by Lutz and Rhoads (1980), Tevesz and Carter
(1980), and Day (1984). According to the hypothesis advanced
by Lutz and Rhoades (1977) from research on marine
molluscs, under conditions favorable to growth, bivalves add
to their shells by the deposition of successive laminae of
calcium carbonate and conchiolin, an organic-rich substance
secreted by the mantle. Periods unfavorable for growth, such
as winter in temperate regions, apparently produce changes
associated with anaerobic metabolism that lead to the deposi-
tion of a thin, dark, organic-rich growth band in the valves.
Conversely, the hypothesis presented by Coker et a/. (1921)
and summarized by Tevesz and Carter (1980) was developed
through research on freshwater mussels. This hypothesis
describes the ‘‘doubling-up”’ of shell layers resulting from
mantle retraction and re-extension which produces the visi-
ble appearance of a dark ring on the shell. Hence, dark an-
nual rings would be produced by the frequent ‘‘doubling-up”’
of the shell along growth edges produced by frequent growth
interruptions from the onset or outset of cold weather (winter).
Either of these hypotheses could explain the prominent an-
nual rings that we observed, formed in winter and visible by
late spring in Virginia.

There was no indication of long-term tagging or mark-
ing stress on shell growth of species recovered for age valida-
tion. Unmarked, freshly dead specimens and shells from
muskrat middens showed growth increments and rates similar
to those in tagged and marked shells of comparable ages
(Moyer, 1984). Brousseau (1979) also reported no significant
differences in growth rate between handled and unhandled
softshell clams (Mya arenaria L.) Handling stress was re-
ported in earlier studies with freshwater mussels (Isley, 1914;
Coker et al., 1921; Negus, 1966), and notching of bivalves can
result in the formation of disturbance lines in shells (Lutz and
Rhoads, 1980). Our handling and marking procedures pro-
bably resulted in some stress of mussels, and disturbance
lines were formed on many specimens that we marked and
later examined. These lines were less prominent than annuli
and apparently were formed at the time of marking. However,
there was no evidence, based on mussel behavior after mark-
ing in the laboratory and comparative growth between marked
and unmarked specimens, that the stress was more than
temporary.

Shell ashing and acetate peels, by the methods
described, proved to be ineffective techniques for use on
freshwater mussels. However, the combination of 5% HCl
etching solution and 15-45 sec etching time provided some
peels of suitable quality. Recent modifications and im-
provements in the acetate peel technique could now make
this method more applicable to freshwater bivalves (Ropes,
1987), and further testing is warranted.

Thin-sectioning of shells was judged to be the most
consistent and accurate technique for age determinations.
Thin-sections provided the highest degree of resolution for
all species examined, and for all sizes and ages, from 15 to
210 mm and 3 to 56 years. Annulus formation was readily ap-
parent in cross-sections of marked shells, and true and false

annuli could be easily separated. Minor shortcomings of the
thin-sectioning technique were the 0.5 to 1 hr required to
prepare a specimen for examination, the need for several cuts
on large shells to fit the petrographic slides (27 x 46 mm) used
in this study, and the difficulty in sectioning small shells
(<20 mm). Because small, thin shells often were too brittle
to withstand the pressure of the cutting blade or chuck used
to hold the shell in place, we suggest that bioplastics be used
for embedding the shells. Modification of the equipment or
technique should overcome these minor problems.

We observed occasional inclusion of small particles
of sediment in shells, which produced the formation of a thick,
dark line internally and externally, especially on female Villosa
vanuxemi, as noted previously. This band was a false annulus
because it was incomplete and usually occurred only in the
vicinity of the foreign particle. Its formation is perhaps
evidence of the adventitious conchiolin layering reported by
Beedham (1965) and reviewed by Tevesz and Carter (1980).
Such layers are described as being a conchiolin-rich damage
response mechanism, often found in unionids having thin-
shelled umbonal areas. They apparently are produced to
mitigate damage caused by extraneous water, sediment, or
other material entering through an abnormal separation be-
tween the mantle and shell margin.

Our test of the growth ring method confirmed the in-
adequacy of this technique, as previously noted by Rhoads
and Lutz (1980). Erosion and corrosion of shells, separation
of true from false annuli, and difficulty in counting closely
deposited growth bands in older shells produced consistent
underestimates of specimen ages. These errors in age, even
on shells with relatively clear annuli such as those of
Fusconaia cor and Pleurobema oviforme, would undoubtedly
occur with most other unionids and result in erroneous ages
and, consequently, imprecise population statistics. Jones et
al. (1978) cautioned that growth curves based on external
growth lines probably underestimate growth rate in young
clams and overestimate it in old ones. Our results with
freshwater mussel shells support this conclusion and indicate
that the growth ring method provides only an estimate of
mussel ages at best, particularly for older cohorts. With the
current availability of sectioning techniques to provide more
accurate ages of unionids, we recommend that use of the
growth ring method be discontinued for all but the younger
age classes or rapidly growing species that are age-validated.

ACKNOWLEDGMENTS

The Virginia Cooperative Fish and Wildlife Research Unit is
jointly supported by the United States Fish and Wildlife Service, the
Virginia Department of Game and Inland Fisheries, Wildlife Manage-
ment Institute, and Virginia Polytechnic Institute and State University.

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INTRACAPSULAR DEVELOPMENT OF THAIS HAEMASTOMA
CANALICULATA (GRAY) (PROSOBRANCHIA: MURICIDAE)
UNDER LABORATORY CONDITIONS

RICHARD A. ROLLER’ AND WILLIAM B. STICKLE
DEPARTMENT OF ZOOLOGY AND PHYSIOLOGY
LOUISIANA STATE UNIVERSITY
BATON ROUGE, LOUISIANA 70803, U. S. A.

ABSTRACT

Copulation and egg capsule deposition of Thais haemastoma canaliculata (Gray) and subse-
quent development of embryos to hatching were investigated. Adult 7’ haemastoma canaliculata deposited
egg capsules, each containing approximately 3200 fertilized eggs. The number of capsules deposited
by any one snail over several days varied between 20-30. The expected ontogeny of spiralean cleavage
followed by gastrula, trochophore, and veliger larva occurred. The trochophore and veliger stages were
easily distinguished from each other. No nurse eggs occur in this species. Hatching of planktotrophic
veligers occurred within 13 days after capsule deposition at 25%9S and 25-26°C. Capsule wall dry
weight decreased significantly; whereas, capsule content dry weight increased during the intracap-
sular period, largely due to increased calcification of embryonic shells. Embryonic calcium levels in-
creased 24 fold during the intracapsular period.

The Southern Oyster Drill Thais haemastoma canali-
culata (Gray) (=7. haysae, Clench, 1927) (Abbott, 1974), is a
muricid gastropod inhabiting estuaries along the Louisiana
gulf coast. This species is the primary predator on the Eastern
Oyster Crassostrea virginica (Gmelin), the only commercial-
ly important species of oyster in Louisiana. It is believed that
T. haemastoma canaliculata represents the greatest hazard
to the survival of C. virginica (Pollard, 1973), thus making the
drill an economically important destructive agent to the oyster
fisheries in Louisiana (St. Amant, 1938, 1957; Burkenroad,
1931). In recent years, salt water intrusions, caused by the
dredging of the Mississippi River at the Gulf of Mexico, have
allowed T. haemastoma canaliculata to migrate further into the
oyster seed grounds thus reducing the economic feasibility
of extensive oyster culture (Pollard, 1973; Van Sickle et a/.,
1976; Smith, 1983). The predation of 7 haemastoma
canaliculata on oysters and the regenerative ability of its
feeding mechanism in response to injury have been previously
described (Garton and Stickle, 1980; Roller et a/., 1984).
Seasonal changes in the reproductive component weights of
the southern oyster drill indicate major episodes of capsule
deposition occurring between April and August (Belisle and
Stickle, 1978).

1Present address: Department of Biology, University of Wisconsin,
Stevens Point, Wisconsin 54481, U.S.A.

Considerable interest in the reproductive biology and
embryology of prosobranch gastropods has stimulated
research by various investigators for many years. These in-
vestigations have varied from complete descriptions of the
embryological development of certain gastropods (Conklin,
1897; Pelseneer, 1911; D’Asaro, 1966) to descriptions of specific
morphological and ecological relationships of various larval
forms (Thorson, 1950; Mileikovsky, 1971; Fretter, 1972; Spight,
1977; Strathmann, 1980; Hadfield, 1984; Pechenik, 1984). St.
St. Amant (1938) provided a well written account of the general
biology of Thais floridana haysae (Clench) (= T. haemastoma
canaliculata); however, very few figures were included in the
work, and the thesis was never published. D’Asaro (1966), us-
ing light microscopy, gave an excellent discussion of the em-
bryogenesis of Thais haemastoma floridana (Conrad). Belisle
and Byrd (1980) used electron microscopy to investigate in
vitro egg activation and development through hatching in
Thais haemastoma. No investigation to date has attempted
to combine the use of light and scanning electron microscopy
(SEM) to view the copulation, ovipositioning, capsule struc-
ture, and developmental stages of 7.) haemastoma canali-
culata. Furthermore, intracapsular weight changes prior to
hatching have not been investigated. Knowledge of embryonic
weight changes prior to hatching would yield valuable infor-
mation concerning possible nutritive contributions of intracap-
sular components.

American Malacological Bulletin, Vol. 6(2) (1988):189-197

189

190 AMER. MALAC. BULL. 6(2) (1988)

While considerable ambiguity exists concerning the ex-
act taxonomic position and classification of Thais spp. of the
Gulf of Mexico (Butler, 1985), the species examined in the pre-
sent investigation was identified as 7) haemastoma canali-
culata (Gray) based on the presence of a large nodular shell
possessing a strongly indented suture (Abbott, 1974). The ob-
jectives of the present investigation were to (1) observe copula-
tion and capsule deposition of adult Thais haemastoma
canaliculata in the laboratory; (2) determine the intracapsular
developmental rate of embryos to hatching at a salinity (25/99)
and temperature (25°C) similar to that experienced in the
estuary; (3) examine changes in capsule structure and com-
position during development; and (4) rear hatched veligers.

MATERIALS AND METHODS

Adult Thais haemastoma canaliculata (shell length
> 40 mm) were collected monthly during 1982 and 1983 from
Bay Champagne near Grand Isle, Louisiana, U.S.A. Snails
were transported to the laboratory and placed into 38 / aquaria
(30 snails/aquarium) containing artificial seawater (Instant
Ocean® Sea Water Mix) at the temperature and salinity of
the collection site (at time of collection). The seawater near
Grand Isle fluctuates in salinity and temperature between 10
and 35% 9 and 10 and 30°C, respectively, over the course of
a year (Barrett, 1971); however, the aquaria were maintained
at constant salinity and temperature during this investigation.
The male:female ratio in each aquarium was approximately
1:1. The snails were maintained on a photoperiod similar to
the natural conditions under which they were collected. Drills
were fed oysters (Crassostrea virginica) and clams [Rangia
cuneata (Sowerby)].

Copulation and capsule deposition in the aquaria were
observed and photographically recorded. Capsules were
removed from the aquaria as soon as possible. Since the cap-
sules were covered by the foot of the snail during deposition
it was often necessary to delay their removal from the aquaria
for several hours.

Individual egg capsules of known age were transferred
to separate, clean glass culture bowls (10 cm tall x 19 cm
diameter) containing filtered (0.45 um) seawater at the ap-
propriate temperature and salinity. The seawater in each bowl
was aerated and changed daily throughout the experiment.
Five capsules were sampled daily for the determination of
developmental rates. lridectomy scissors were used to open
the egg capsules. The embryos were removed with a pasteur
pipet and placed on glass slides with clay-supported
coverslips. Embryos were then examined and photographed
with a Leitz Wetzlar Orthoplan compound microscope with
an Orthomat camera attachment. Embryos obtained from in-
dividual capsules were examined to determine if development
to hatching was synchronous within a particular capsule. In-
tact and opened capsules were photographed with a Wild TYP
stereo-dissection microscope with a Nikon M35-S camera at-
tachment. Intracapsular osmolarity was determined with a
Wescor vapor pressure osmometer.

Two days after deposition, ten capsules were opened
and the embryos were removed and counted. Approximately

one day prior to hatching, 10 randomly selected capsules from
each culture bowl were dissected for mortality determination.
Each culture bowl was examined daily for hatched
veligers, which were then transferred to additional culture
bowls containing freshly aerated and filtered sea water
(1 larva/100 ml). The water in each bowl was replaced daily.
Veligers were then fed 104 cells/ml (final concentration) daily
of /sochrysis galbana (Parke) - Monochrysis lutheri (Droop)
(1:1). Algae were cultured using the method of Guilliard (1975).
For scanning electron microscopy (SEM), embryos
were removed from the capsules for fixation. Veligers were
first anesthetized with MgSO, and then fixed for SEM. The
best anesthetization was achieved by slowly adding small
amounts (approximately 0.19) of granular MgSO, to the culture
water until the larvae were completely immobile but had not
contracted or withdrawn into their shells. Specimens were
fixed overnight with 2.5% gluteraldehyde in 0.2M sodium
cacodylate-sucrose buffer (731 mOsm; pH = 80). The
sucrose was used to adjust the osmolality of the fixative to
the appropriate salinity of the culture in order to reduce
osmotic stress during fixation. After fixation, the specimens
were rinsed in three changes of distilled water to remove all
buffer salts, dehydrated in acidified 2,2-dimethoxypropane
(DMP), and transferred to modified Beem™ capsules with a
25 um Nitex screen over each end. The specimens were then
critical-point dried in CO2, coated with approximately 200 A
of Au/Pd, and examined with a Hitachi S-500 scanning elec-
tron microscope at 25 KV. Empty egg capsules were sectioned
with a razor blade and prepared as above for SEM investiga-
tion. For light microscopy, intact capsules containing embryos
and larvae were fixed overnight in formalin-acetic acid-alcohol
(FAA), dehydrated in ethanol, cleared with xylene, embedd-
ed in paraffin, sectioned at 7 um, and stained with Azan
(Humason, 1972).
For capsule dry weight analysis, random samples of
20 capsules were taken one day after deposition (Day 1) and
three days prior to hatching (Day 10). The total length of each
capsule was measured with a vernier caliper. Each capsule
was briefly rinsed in distilled water and then dissected into
two components: capsule wall and capsule contents (embryos
and albumen). The components were then lyophylized and
capsule wall dry weight, capsule content dry weight, and total
capsule dry weight was determined to 0.001 mg using an
analytical balance. Capsule component indices were then
calculated by the method of Stickle (1973). The relationship
between capsule length and dry weight was analyzed by sim-
ple linear regression (SAS Institute Inc., 1985a, b). Differences
between Day 1 and Day 10 dry weight components were com-
pared by a two-sample t-test (Steel and Torrie, 1980).
Embryonic calcium levels were analyzed by atomic ab-
sorption spectrophotometry (Perkin-Elmer Corp., 1982). Twen-
ty capsules on Day 1 and Day 10 were dissected and the con-
tents were incubated in 10 ml of a 1% LaO3 / 5% HCI mixture
(40°C) for 1 hour to mobilize any calcium present. The con-
tents of each capsule were then centrifuged. The superna-
tant was removed, diluted 2X with fresh LaO3-HCl, and ana-
lyzed. Total inorganic material was determined on an addi-
tional sample of 20 capsules by ashing at 450°C for 4 hours.

ROLLER AND STICKLE: THAIS INTRACAPSULAR DEVELOPMENT 191

Total organic material was calculated by subtracting the total
ash (inorganic) from the pre-combustion dry weight. Day 1
and Day 10 calcium, organic, and other inorganic levels were
compared by a two-sample t-test (Steel and Torrie, 1980).

RESULTS

COPULATION AND CAPSULE DEPOSITION

Copulation in the drills was observed in the field from
late April to late June, 1982 and from late April to early June,
1983. During these months snails were found in large breeding
aggregations which extended from approximately 0.5 m above
the water surface at low tide to 1 m in depths. The number
of snails comprising each aggregation varied from 6 to 27 in-
dividuals. Drills collected in early June, 1983 began copulating
in the laboratory within 5 days. The duration of copulation was
variable, lasting from approximately 2.5 hours to 3 days. Dur-
ing copulation the male crawled onto the shell of its partner
and inserted its penis into the right side of the mantle cavity.
Spermatozoa and prostatic secretions were presumably dis-
charged into the genital aperture of the female (Fretter and
Graham, 1962).

Egg capsule deposition occurred as early as six hours
and up to sixty days after copulation was observed. In the
laboratory, the egg capsules were attached to the glass walls
of the aquaria, usually near the exhalant port of the
undergravel filter system. Rarely were capsules deposited on
oyster shells; however, oysters covered with Thais egg cap-
sules have been collected from Grand Isle. Capsule deposi-
tion was intermittent. Snails were observed to cease deposi-
tion for a while, feed on oysters, and then resume deposition,
sometimes in an entirely different location. Snails tended to
attach their capsules together forming one large communal
mass. The intermittent feeding behavior as described above
and the communal egg masses made distinguishing which
female laid specific capsules difficult. The number of capsules
obtained from any one snail varied; however, most drills
deposited 20-30 capsules in a mass. The duration of capsule
deposition also varied, from as short as 2-3 hours to as long
as 6-7 days. Snails were also observed to pause during
deposition and remain on the capsule mass without feeding
for several hours before resuming capsule laying.

Capsules were usually attached by their bases (Fig.
1), and formed a single layer on the substratum. In several
cases capsules were observed attached together at various
locations along their lengths; however, attachment never
obstructed the opercular opening of any capsule in a mass.
Butler (1954) reported similar findings.

The egg capsules of Thais haemastoma canaliculata
are similar to those of 7 haemastoma floridana as described
by D’Asaro (1966). The capsules are somewhat conical in ap-
pearance, possessing a broad flat apical plate and tapering
down to the base where they are typically attached to the
substratum (Fig. 1). Each capsule possesses a convex and
concave side along most of its length, giving the capsule an
oblong appearance in cross section at the distal end (Fig. 2).
However, the capsule is more circular in cross-section at its

Fig. 1. Light micrograph of typical Thais haemastoma egg cases con-
taining embryos. Hatching occurred approximately three days later
(O, operculum). Fig. 2. Scanning electron micrograph (SEM) of an
opercular view of an egg capsule (Cc, concave wall; Cv, convex wall;
O, opercular plug; P, one lateral protuberance). Fig. 3. SEM of cap-
sule cross-section showing both inner and outer walls (lw, inner cap-
sule wall; Ow, outer capsule wall; P, lateral protuberance). Fig. 4.
SEM of capsule protuberance outlined in (3) (D, lateral dense layers
of outer capsule wall; Iw, inner capsule wall; S, medial spongy mass
of outer capsule wall).

tapered base. Four longitudinal ridges (2 on each side)
separate the convex and concave sides. The two ridges on
each side merge at the apical plate forming a lateral pro-
tuberance (Figs. 2-4). Each capsule is composed of a thick
fibrous-appearing outer wall and a thin membranous inner
wall, which readily separate during microscopical preparation
(Fig. 3). The entire outer capsule wall appears to be composed
of two compact, dense lateral layers and a spongy-fibrous
medial layer (Fig. 4). The protuberances and ridges repre-
sent sculpturing of the outer wall only and do not make up
any portion of the inner wall, which encloses the embryos and
the nutritive albumen. A round, discoidal opercular plug is
located on the apical plate at the distal end of each capsule
(Fig. 2). The operculum swells and bulges outward a few days
prior to hatching. At hatching the operculum disintegrates
leaving a prominent opercular scar.

Capsule length varied from 0.84-1.13 cm (x + S.E.=
0.95 + 0.01 cm; N=40). Capsule wall and capsule content
dry weight varied from 0.47-1.57 mg (x + S.E.= 1.05 + 0.05
mg; N=40) and from 0.14-1.20 mg (x + S.E.= 0.54 + 0.04

192 AMER. MALAC. BULL. 6(2) (1988)

mg; N=40) respectively. The total capsule dry weight varied
from 0.92-2.14 mg (x + S.E.= 1.60 + 0.06 mg; N=40). Cap-
sule wall and content dry weight comprised 65.6 + 2.3 and
34.4 + 2.3(x + S.E.) percent, respectively, of the total cap-
sule dry weight. Capsule wall dry weight varied directly with
capsule length: dry weight (mg)= -2.54 + (3.79 X length in
cm) (r2=0.673; N=40; P<0.001). A significant linear regres-
sion of total capsule dry weight on length also existed and
is given as dry weight (mg)= -1.48 + (3.28 X length in cm)
(r2=0.468; N=40; P< 0.001). No significant relationship existed
between capsule content dry weight and capsule length
(P>0.05). Each capsule contained 3246 + 21 (x + S.E.;
N=10) embryos embedded in a viscous, albuminous fluid.
Capsules, when deposited, were a milky white color, which
during development turned light tan and finally dark brown
just prior to hatching. Only three capsules deposited in the
laboratory developed the dark purple color, characteristic of
dead or stressed embryos (St. Amant, 1938; D’Asaro, 1966;
Spight, 1977; Pechenik, 1982; Butler, 1954, 1985). Examina-
tion of these capsules revealed that all embryos were dead.

DEVELOPMENTAL RATE AND STAGES

Development of Thais haemastoma canaliculata was
synchronous within a particular capsule throughout the en-
tire period of encapsulation and required 12-13 days to hatch-
ing at 25% 9S and 25°C (Table 1). Unfertilized eggs were
spherical and approximately 65-70 um in diameter; however,
as reported previously (St. Amant, 1938; D’Asaro, 1966), the
majority of the yolk (deutoplasm) was concentrated in one pole
(vegetal) with other cytoplasmic constituents being concen-
trated at the opposite (animal) pole. First and second polar
body formation was complete within 2.5 hours after deposi-
tion of the capsule. By the second polar body stage (Fig. 5),
the fertilized egg had elongated and the animal and vegetal
areas were easily distinguished. The round yolk granules in
the vegetal area were visible in live and preserved (Figs. 5,
6) zygotes. Early cleavage was restricted to the animal pole
of the embryo. The first cleavage, producing the AB and CD
blastomeres (Fig. 7) occurred 5-6 hours after deposition (Table
1). The second cleavage (Fig. 8) occurred within 2-4 hours
after the first cleavage. As D’Asaro (1966) showed for 7.

Table 1. Developmental rate of Thais haemastoma canaliculata at
25°%o9S and 25-26°C.

Developmental Event Time
Fertilized egg with 2 polar bodies 2.5 hours
First cleavage 5-6 hours
Second cleavage 8-9 hours
16 cell stage 17-19 hours
Stereoblastula 28 hours
Early gastrula 3.5-4 days
Stomodael invagination, cephalic

expansion & shell gland formation 5 days
Trochophore 5.5-6 days
Early veliger 7 days
Hatching 13 days

haemastoma floridana, we found that the D blastomere
possessed a large polar lobe (Fig. 8). Within 17-19 hours after
capsule deposition, the 18 cell stage was complete. By that
time, the polar lobe had been resorbed, and the large 2D
macromere was seen (Fig. 9).

A stereoblastula containing a narrow segmentation
cavity, as reported by St. Amant (1938), formed approximate-
ly 9-11 hours after polar lobe resorption (Table 1). Gastrula-
tion by epiboly and archenteron formation (Fig. 10) was

Fig. 5. SEM of fertilized egg after second polar body formation (Cy,
cytoplasmic (animal) pole; Pb, polar bodies; V, vegetal yolk-containing
pole). Fig. 6. SEM of vegetal view of ruptured polar lobe, illustrating
dense yolk mass (y, yolk mass). Fig. 7. SEM showing first cleavage
of the ovum, resulting in formation of AB and CD cells (Pb, polar
bodies; Pl, polar lobe). Fig. 8. SEM of four-cell stage showing com-
pletion of A, B, C, and D cells with polar lobe (Pl) and polar bodies
(Pb) still evident. Fig. 9. SEM of 2D cell, after polar lobe resorption
(Pb, polar bodies). Fig. 10. SEM of gastrula stage, illustrating the
blastopore (BI).

ROLLER AND STICKLE: THA/S INTRACAPSULAR DEVELOPMENT 193

observed within 3.5-4 days after oviposition. Stomodaeal in-
vagination, cephalic expansion, and formation of the shell field
invagination (Fig. 11) occurred 5 days after deposition and
followed the same pattern as described for Thais haemastoma
floridana (D’Asaro, 1966).

The early trochophore (Fig. 12) was characterized by
a prominent stomodaeum, an apical tuft, the beginning of pro-
totrochal and telotrochal ciliation, and the appearance of the
larval kidneys. The late trochophore stage (Fig. 13) exhibited
antero-posterior elongation, prominent larval kidneys, and well
formed prototrochal, metatrochal, and telotrochal ciliation. The
early veliger stage was characterized by the presence of the
velar ciliation (Fig. 14). The dorsal margin of the shell gland
was complete, and the protoconch covered the posterior
region of the digestive gland’s primordial cells. At this stage,
the operculum was first evident (Fig. 15). By 8 days after cap-
sule deposition, torsion, which results in a 180° rotation of the
visceral mass, was complete. At this time, the apical ciliation
and operculum were well developed, and the ventral foot and
larval tentacles were first seen (Figs. 16-18).

No nurse eggs, as described by Rivest (1983), were
observed. The viscosity of the intracapsular contents declined
over the course of the developmental period; however, the
measured intracapsular osmolarity did not change during
development. It is therefore possible that the intracapsular
albumen is consumed by the embryos and replaced by sea
water.

CAPSULAR CONTENT CHANGES DURING
DEVELOPMENT

Capsule weight changes prior to hatching are il-
lustrated in Table 2. During the intracapsular developmental
period, the weight of the capsule contents significantly in-
creased 63.0%; capsule wall weight decreased 43.4%; and
the total capsule weight (contents and wall) decreased 18.4%.
Total capsule ash significantly increased 37.8%, while total
capsule calcium increased 24-fold over the encapsulated
developmental period. Total capsule organic material
significantly decreased 37.7%; however, other inorganic
material (excluding calcium) showed a non-significant in-
crease of 2.0%.

HATCHING AND REARING OF VELIGERS

Hatching of veligers (Figs. 17, 18) at 25%/p9S and 25°C
occurred between 12-13 days after capsule deposition. The
shell length at hatching was 49.7 + 8.3 um. Hatching was ac-
complished through the dissolution of the capsule’s oper-
culum, possibly by mechanical means (St. Amant, 1938) or
by chemical means (Sullivan and Bonar, 1984). Most
(96-100%) embryos developed into normal appearing veligers
and survived to hatching. In some capsules approximately
2-4% of the veligers were either dead or malformed at hatch-
ing. Hatched veliger larvae survived up to 50-53 days when
kept in laboratory cultures and fed a mixture of /sochrysis
galbana and Monochrysis lutheri. Ninety percent of the
hatched veligers survived 45-50 days in culture. The shell

fe Si)

Fig. 11. Light micrograph showing stomodael invagination (S) and
apical plate formation (Ap), immediately after gastrulation and prior
to formation of trochophore (D, digestive system primordium; Sg, shell
gland). Fig. 12. SEM of early trochophore stage (At, Apical tuft cilia-
tion; Lk, larval kidney; Pt, prototrochal ciliation; S, stomodaeum; Tt,
telotrochal ciliation). Fig. 13. SEM of late trochophore stage, show-
ing formation of metatrochal ciliation (Lk, larval kidney; Mt, metatrochal
ciliation; Pt, prototrochal ciliation; S, stomodaeum; Tt, telotrochal cilia-
tion). Fig. 14. SEM of a dorsolateral view of early veliger larva (A,
Apical ciliation; Lk, larval kidneys; P, protoconch; V, velum). Fig. 15.
SEM illustrating a ventral view of an early veliger larva, illustrating
shell operculum formation and prominent shell gland ciliation (Lk,
larval kidneys; Op, shell operculum; P, protoconch; Sg, shell gland;
V, velum). Fig. 16. Light micrograph illustrating a midsaggital sec-
tion (7 um) through a veliger (three days prior to hatching), showing
further elongation of foot and operculum (F, foot; Op, shell operculum;
P, protoconch; V, velum).

length of the veligers after 37 days in culture was 122.4 +
28.3 «pm. No settlement/metamorphosis occurred, even though
the larvae appeared healthy and fed on the algal species pro-
vided (Fig. 18).

194 AMER. MALAC. BULL. 6(2) (1988)

Table 2. Capsule component weights on Day 1 and Day 10 for Thais
haemastoma canaliculata capsules. Capsule components (in mg) are
separated into organic, Ca?*, and other inorganic components. N=
40 capsules.

DAY DAY T
1 10 VALUE
CAPSULE CONTENTS
Organics 0.090 + 0.005* 0.181 + 0.004 14.844
Calcium 560 x 104 + 5.2 x 10°6 0.161 + 0.003 55.46t
Other
Inorganics 0.323 + 0.012 0.332 + 0.009 0.56 N.S
Total 0.414 + 0.044 0.675 + 0.056 3.644
CAPSULE WALL
Organics 1.218 + 0.031 0.634 + 0.039 11.694
Calcium 6.41 x 10°3 + 1.6 x 10-4 7.21 x 103 + 4.14+
1.0 x 10-4
Other
Inorganics 0.119 + 0.004 0.119 + 0,004 0.03 N.S
Total 1.344 + 0.033 0.760 + 0.038 11.614
CAPSULE TOTAL (wall and contents)
Organics 1.309 + 0.056 0.815 + 0.077 5.18t
Calcium 6.97 x 10°3 + 89 x 10°5 0.168 + 0.003 54.46t
Other
Inorganics 0.443 + 0.012 0.452 + 0.010 0.56 N.S.
Total 1.759 + 0.054 1.435 + 0.001 3.29T
*.-Mean + SE.
$ - Statistically significant at « =0.001
t - Statistically significant at « =0.01
N.S. - Nonsignificant
DISCUSSION

We observed, with the aid of scanning electron
microscopy, a distinct trochophore stage (Figs. 12, 13) for Thais
haemastoma canaliculata. St. Amant (1938) had earlier
reported that in 7. floridana haysae the trochophore was
atrochal and could not be distinguished from the early veliger
stage; therefore, the early veliger could be identified only after
the shell was formed. The development of the velum (Fretter
and Graham, 1962), which is difficult to observe using stan-
dard light microscopy (St. Amant, 1938), is easily seen using
SEM techniques (Figs. 12-15). The development of the pro-
toconch is also more apparent from SEM observations (Fig.
14). St. Amant was unable to identify the onset of torsion in
this species. We found that in ZT haemastoma canaliculata tor-
sion occurs prior to hatching, which agrees with D’Asaro’s
(1966) observations of 7. haemastoma floridana development.

Belisle and Byrd (1980) identified two different cleavage
patterns occurring in Thais haemastoma, as opposed to the
one distinct pattern reported by St. Amant (1938) and D’Asaro
(1966). In the present study, we failed to observe the second
cleavage pattern observed by Belisle and Byrd (1980). The
only cleavage pattern we observed agrees with that reported
by D’Asaro (1966). The absence of the second cleavage pat-
tern in our investigation does not deny its existence. This
second pattern could be an infrequent deviation from the “‘nor-
mal’’ pattern usually observed.

18

Fig. 17. SEM showing an anterior view of a hatched veliger, illustrating
a well developed velum (V), larval tentacle (T), and cephalic ciliation
(C) (F, foot; M, mouth; Op, shell operculum). Fig. 18. Light micrograph
of newly hatched veligers, illustrating prominent structures (A, algal
cell in gut; Ag, anal gland; F, foot; |, intestine; Op, shell operculum;
S, stomach; V, velar cilia).

The difference in developmental rate observed in the
present study (13 days to hatching at 25%o9S and 25-26°C)
and that observed by Belisle and Byrd (1980) (16 days to
hatching at 207/99 and 24°C) could be due to differences in
experimental temperature and salinity. Belisle and Byrd (1980)
did not specify the subspecies of snail they studied. The
developmental patterns and rates we observed for Thais
haemastoma canaliculata at 25%9S and 25-26°C are very
similar to those reported for 7. haemastoma floridana by
D’Asaro (1966). The ranges of these two subspecies overlap
and both are found on the Louisiana coast, although 7.
haemastoma canaliculata is more numerous (St. Amant,
1938). Butler (1954) made reciprocal crosses between the two
subspecies and obtained normal larval development, sug-
gesting that hybridization could occur in this area. Since the
embryology of 7. haemastoma canaliculata and T.
haemastoma floridana is similar (St. Amant, 1938; Butler, 1954;
D’Asaro, 1966; present study), the separation of the two into
separate subspecies based on shell morphology alone is
possibly unjustified. Further data, in the form of electrophoretic
analysis, are needed.

Hatching of veligers, in the present study, occurred be-
tween 12-13 days after oviposition and was possibly accom-
plished by chemical dissolution of the capsule operculum,
as occurs in the mud Snail //vanassa obsoleta (Say) (Sullivan
and Bonar, 1984). Veligers in laboratory culture survived 50-53
days after hatching but did not metamorphose. Algal cells
were observed in the gut of the veligers (Fig. 18) and the lar-
vae appeared healthy; however, none survived to settlement
and metamorphosis. It is possible that the veligers did not
obtain enough nutrients from the algal cells provided. The
veligers did survive an extended time (50 days) and showed
evidence of some growth (from 49.7 um to 122.4 um). Further-
more, the algal species and concentration provided have been
sufficient for other planktotrophic larvae (Ament, 1979;
Jespersen and Olsen, 1982; Sprung, 1984); however, the
nutrient levels and quality necessary for maintenance could
possibly not be sufficient for growth and metamorphosis. Had-
field (1984) showed that a high degree of substratum chemical

ROLLER AND STICKLE: THA/S INTRACAPSULAR DEVELOPMENT 195

specificity can be required to induce settlement and metamor-
phosis in molluscan larvae. Several chemical compounds,
from naturally occurring substances, have been identified as
inducers of larval settlement and metamorphosis (Morse et
al., 1979; Heslinga, 1981; Rumrill and Cameron, 1983; Morse
and Morse, 1984). It is possible that one or more inducers
exist for Thais haemastoma canaliculata. Such inducers could
exist in encrusting algae on oyster shells or possibly in
polychaete tubes or barnacles upon which young Thais prey.
Oysters, tube-dwelling polychaetes, and barnacles are abun-
dant along the Louisiana coast.

Gastropod species possessing teleplanic veligers
could be dispersed over a large geographic range and would
have a planktonic existance of long duration (Scheltema,
1978). It appears from our results and the observations of
others (St. Amant, 1938; D’Asaro, 1966; Scheltema, 1978) that
Thais haemastoma canaliculata veligers are teleplanic and
are likely to survive as long in the field as they did in the
laboratory.

The spongy/dense layering of the outer wall of the egg
capsules could just be the result of the process used to form
the capsule and have no specific function; however, in our
opinion this layering appears similar to that seen in vertebrate
long bones (Mader, 1985) and could possibly aid in lending
strength and support to the capsules thus protecting the
enclosed embryos against physical damage. The protuber-
ances and ridges could aid in maintaining an upright cap-
sule and further enhance the protection of the delicate em-
bryos inside.

Egg capsule dry weight varied directly with capsule
length (r2= 0.673; P<0.001) for Thais haemastoma
canaliculata. The dry weight of individual capsule components
varied differently from ovipositioning to hatching. The overall
decrease in total capsule dry weight during the intracapsular
developmental period (Table 2), possibly reflects the loss of
metabolic end products through the capsule wall. The weight
of a single capsule operculum (unpublished data) is only 0.08
+ 0.02 mg (N = 20); therefore, the 43.4% decrease in cap-
sule wall weight appears too high to be explained solely by
chemical dissolution of the opercular cap. It is possible that
portions of the inner matrix of the capsule wall are eroded
prior to hatching; however, in this investigation all hatched
veligers exited from the capsule through the operculum. It
is therefore unlikely that erosion of other portions of the cap-
sule wall would aid the hatching process by forming multiple
exits. It is possible that nutrients or other substances are
removed from the wall and utilized by the developing embryos.
Since the majority of the capsule wall is composed of organic
material (Table 2) and the uptake of dissolved organic material
(DOM) by molluscan larvae has been documented (Manahan,
1983), this hypothesis is possible. This hypothesis could also
aid in explaining the doubling of the capsule content organic
weight; however, this is speculative since we have no confirm-
ing data. The dry weight of the capsule contents (albumen
and embryos) significantly increased 63% prior to hatching.
We have shown (Table 2) that much of this increase (24%)
is due to the uptake of calcium by the embryos, presumably
for calcification of the shell prior to hatching. Eyster (1986)

showed that calcium was the main constituent of early shell
mineralization for several species of gastropod veligers.
Likewise, our data for T haemastoma canaliculata support an
observed overall increase in calcium content of these veligers
prior to hatching. We found a small amount of calcium
associated with the capsule wall (Table 2), which was probably
due to residual calcium adsorbtion to the wall, or possibly a
small number of embryos that we neglected to remove. None
of the increase in capsular content dry weight (i.e. embryonic
weight) was due to inorganic materials other than calcium.
It is expected of marine organisms with planktotrophic larvae
that most of the organic growth should occur after hatching
and during the planktonic existence (Scheltema, 1967; Pilk-
ington and Fretter, 1970; Pechenik and Fisher, 1979; Pechenik,
1980, 1984; Pechenik and Lima, 1984). We observed a signifi-
cant doubling in the organic material of the capsule contents;
however, we made no weight measurements of planktonic
veligers. The increase in the organic material of the capsule
contents could be related to the corresponding loss of organic
material from the capsule walls; however we have no data
to prove this assumption. It is clear that observations on
growth and weight changes of planktonic stages is needed
before any comparisons can be attempted.

It was the purpose of this study to add to earlier in-
vestigations of Thais developmental patterns, egg capsule
structure, and weight changes over the course of intracap-
sular development. Our findings and those of St. Amant
(1938), Butler (1954), D’Asaro (1966), and Belisle and Byrd
(1980) illustrate the tremendous reproductive potential for this
species. Even though the planktonic larval mortality must be
quite high, when one considers the sheer number of embryos
contained in a single capsule (about 3200), the number of
capsules deposited by a single female (20-30), and the
96-100% survival to hatching (laboratory conditions), it is no
surprise that this species is a serious economic threat to the
oyster industry along the United States gulf coast.

ACKNOWLEDGMENTS

We thank M. Kapper, J. Lynn, M. Holley, and the anonymous
reviewers for corrections to the manuscript. Appreciation is extend-
ed to Dr. J. Fleeger and Dr. E. Weidner for their advice during the
course of this investigation. Special thanks are expressed to Tina
F. Roller for her assistance. This research was funded in part by a
grant from the Petroleum Refiners Environmental Council of Loui-
siana (PRECOL) and from NSF Grant No. DEB-7921825.

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Date of manuscript acceptance: 26 June 1987

vy

TEMPORAL AND SPATIAL VARIATION OF SHELL MICROSTRUCTURE
OF POLYMESODA CAROLINIANA (BIVALVIA: HETERODONTA)

ANTONIETO TAN TIU'
DEPARTMENT OF BIOLOGICAL SCIENCES
UNIVERSITY OF SOUTHERN MISSISSIPPI

SOUTHERN STATION, BOX 5018
HATTIESBURG, MISSISSIPPI 39406-5018, U. S. A.

ABSTRACT

Temporal and spatial variation of the microstructure of inner surface of shell, condition index
and organic content of shell of the Carolina marsh clam Polymesoda caroliniana (Bosc) in three dif-
ferent Mississippi habitats are described and discussed in relation to one another and environmental
conditions. Microstructure of the inner shell surface distal to the pallial line showed distinct seasonal
variation but little spatial variation. Pseudospiral microstructure, on inner surface of shell undertucked
by the periostracum, predominated over ‘‘normal’’ crossed-lamellar microstructures in cooler seasons.
Presence and seasonal frequency of occurrence of complex crossed-lamella one inside the pallial line
reflected habitat differences. It was consistently present in submerged clams, present only in June
and September in wild clams, and absent in exposed clams. Survival and condition index of transplanted
clams in submerged area were higher than those clams in areas often exposed to air. Condition index
showed seasonal and spatial variation, while organic content did not.

The shell microstructure of a bivalve is determined by
its genome. This genotype sets constraints that fix limits within
which adaptive change can occur. Moreover, while basic
molluscan shell microstructures are few (Taylor et a/., 1969,
1973; Gregoire, 1972; Carter, 1980; Watabe, 1981; Wilbur and
Saleuddin, 1983; Carter and Clark, 1985), subtle variations
within each structural category occur because details of shell
crystallization can be influenced by environmental factors
(Barker, 1964; Taylor et a/., 1969; Rhoads and Panella, 1970;
Lutz and Rhoads, 1978, 1980; Carriker et a/., 1980; Carter,
1980; Prezant and Chalermwat, 1983; Lutz and Clark, 1984;
Carter and Clark, 1985; Prezant and Tan Tiu, 1986; Tan Tiu,
1987; Tan Tiu and Prezant, 1987). Conservative shell micro-
structures can be important characters used to determine
phylogeny (Carter, 1980). Furthermore, consistent, inducible
microstructures could be used to monitor recent or past en-
vironmental conditions (Lutz and Rhoads, 1980). Thus, it is
important to examine shell microstructural variations to
reasonably evaluate the environmental significance of micro-
structural patterns. The goal of this study was to investigate
the extent of shell microstructural variation, temporally and

1Current Address: Harbor Branch Oceanographic Institution, Inc.,
Division of Applied Biology, 5600 Old Dixie Highway, Fort Pierce,
Florida 34946, U.S.A.

spatially, on the eurytopic Carolina marsh clam, Polymesoda
caroliniana Bosc, 1801.

Aragonitic shells of Corbiculidae, like the marsh clam,
consist of outer crossed-lamellar and inner complex crossed-
lamellar layers, separated from each other by a distinct (or
indistinct) myostracum (Taylor et a/., 1973). Shell microstruc-
ture of Polymesoda caroliniana has not been previously ex-
amined in detail except for the conchiolin layers within the
shell (Kat, 1985). Taylor et a/. (1973) briefly described the shell
microstructure of a related species, Polymesoda anomala
(Deshayes, 1855), from Ecuador.

MATERIALS AND METHODS

Specimens of Polymesoda caroliniana, ranging 11 to
43 mm maximum anterior posterior length, were collected
seasonally (June, Sept, Dec 1985, Mar, June 1986) from a
marsh at the Rod and Reel Fishing Camp, Old Fort Bayou,
Jackson County, Mississippi, U.S.A. Each seasonal sample
was treated similarly. Thirty specimens were shucked in the
field. After shell length, height and width were measured,
shells were preserved in absolute ethanol for later examina-
tion by scanning electron microscopy. Areas of inner shell sur-
face examined and compared are shown in figure 1. Another
fifty specimens were transported to the laboratory where

American Malacological Bulletin, Vol. 6(2) (1988):199-206

199

200 AMER. MALAC. BULL. 6(2) (1988)

GS kal 5) Ge

So a

xX a

ae A
Fig. 1. Left valve of Polymesoda caroliniana. Areas of the shell sur-
face examined are marked by dots, corresponding to the letters on
the right (A, area undertucked by periostracum. B, area just dorsal
to Area A. C, area between Area B and pallial line. D, E, F, the ‘‘tran-
sition zone’’. G, area at the level of ventral margin of adductor scars.

H, area at the level of dorsal margin of adductor scars. |, area near
umbo).

length, width, height, total weight with and without mantle
water, shell and tissue dry weight, and organic content of shell
were measured. Condition index and organic content of shell
were computed. Definitions, procedures and care of speci-
mens followed those by Prezant and Tan Tiu (1986) and Tan
Tiu (1987).

A large sample of Polymesoda caroliniana, 11 - 43 mm
long, collected in June 1985 from the Rod and Reel Fishing
Camp, were marked and divided into two groups. One group
was transplanted to a continually submerged area, and the
other to a periodically exposed marsh area. Submerged and
exposed areas are located within a 100 m radius of Halstead
Bayou (adjacent to Gulf Coast Research Laboratory), Ocean
Springs, Jackson County, Mississippi. Each group consisted
of eight cages each containing 45 individuals. Details of pro-
cedures for marking, care of samples, size of cages and how
they were set are similar to those described for studies of Cor-
bicula fluminea Muller, 1774 by Tan Tiu (1987). Two cages were
recovered from each site each season (beginning September
1985) at the same time wild samples were collected from the
Rod and Reel Fishing Camp. Samples were treated as previ-
ously described. Because of high mortality, all cages in the
exposed area were recovered in December 1985.

Monthly measurements of air, ground, surface and bot-
tom water temperatures, water conductivity, dissolved oxygen,
pH, methyl orange alkalinity, total filtrable residue, turbidity,
transparency (Secchi depth), salinity, hardness, calcium, water
depth and organic content of sediment were made in the three
sampling areas. Water was absent in the emerged area on
several occasions (June, July, Nov, and Dec 1985). Thus, water
parameters could not be measured at those times. Details of
methods used and errors of measurement are described in
Tan Tiu (1987).

Significance of seasonal and habitat variation in con-
dition indices and organic content of shell determined by one-
way ANOVA, followed by Tukey test when ANOVA was signifi-
cant. When only two samples were compared, t-test was used.
Subjective evaluation was made in cases where statistical
evaluation was not possible. All statistics were compared with
Critical values at a = 0.05 and critical values used were con-
servative. Statistical methods used are described by Zar
(1984).

Clams collected in the marsh at Rod and Reel Fishing
Camp from June 1985 to June 1986 that were not in cages
nor marked will be referred to as ‘‘wild’’ clams or group. Clams
in cages transplanted to Halstead Bayou will be referred to
as ‘experimental’ groups. Experimental clams that were
placed in the continually submerged area will be referred to
as ‘‘submerged’”’ clams or group, while clams that were placed
in a regularly exposed area will be referred to as ‘‘exposed”’
clams or group.

RESULTS

ENVIRONMENT

The macroflora of the Rod and Reel Fishing Camp
marsh (the location of seasonal wild samples and original
source of experimental samples) and the exposed marsh area
(a transplantation site), is predominantly Juncus roemerianus
Scheele, while the submerged area (another transplantation
site) was devoid of vegetation and had a muddy substratum.
Turbidity, salinity, pH, calcium, total filtrable residue of water
measured in the submerged area were significantly higher
than at the Rod and Reel Fishing Camp (Table 1). During a
few sampling periods (August to October 1985), when water
was present in the exposed area, measurements of turbidity,
conductivity, dissolved oxygen, methyl orange alkalinity and
organic content of sediment were higher in the exposed area
than in the submerged area at the same time. Temperature
of water bottom, as measured by a maximum-minimum ther-
mometer, ranged from 14.0 to 36.0°C in the submerged area
and 7.9 to 42.2°C in the exposed area. Ground temperature
measured at the bank of the submerged area ranged from
7.2 to 42.49°C. No maximum-minimum thermometer data are
available in Rod and Reel Fishing Camp site.

SHELL MICROSTRUCTURE

Condescriptive statistics of the dimensions of shells
examined by scanning electron microscopy are presented in
Table 2. The inner shell surface of Polymesoda caroliniana,
near the umbo (Area |), has irregular pits and grooves. Ven-
tral to Area | (Areas G and H), the microstructures can be
“clumped”’ into irregular mounds (Fig. 2), whose surficial
borders represent areas where lamellae of opposing orienta-
tions meet (Fig. 3). The inner shell surface of areas G and
H may be flattened with few mounds (Fig. 4), or underlain
by irregular to granulate reticulated layers (Fig. 5).

The inner shell surface proximal to the pallial line
(Areas D to I) can be divided into four microstructural types:
complex crossed-lamella one, complex crossed-lamella two,

TAN TIU: POLYMESODA CAROLINIANA SHELL MICROSTRUCTURE 201

Table 1. Condescriptive statistics and t-tests of environmental variables measured at the Rod and Reel Fishing Camp and Submerged Area,
Ocean Springs, Mississippi. Abbreviations in the variable column are as follows: surface water temperature (SWT), turbidity as measured by
a nephelometer (Tur), water transparency as measured by a Secchi disc (Sd), conductivity (Con), dissolved oxygen (DO), salinity (Sal), methyl
orange alkalinity (MOA), calcium (Ca), total filtrable residue (TFR), hardness (Hds) and sediment organic content (SOC). T-statistics of averages
(with one standard deviation and n number of monthly measurements) are evaluated using critical values at a = 0.05 for (df) degrees of freedom.
Min = minimum, max = maximum. Ho: Average values of environmental factors are the same in both places.

Rod and Reel Fishing Camp Submerged Area Significance
Variable range mean standard n range mean standard n computed t critical df
min max deviation min max deviation value

SWT (°C) 195 400 25.9 6.1 13 9.0 31.0 23.4 69 13 Student’s 2.064 24
t = 0976

Tur (NTU) 44 200 8.0 44 12 5.0 26.7 12.9 68 12 Student’s 2.074 22
t = 2.118

Sd (cm) 30 93 66 23 12 35 65 46 11 11 Welch 2.120 16
t = 2.823

Con (umhos/cm) 100 12000 4898 4282 13 750 24000 8142 8901 13 Welch 2.110 AT,
t = 1.184

DO (mg/L) 0 10.0 59 24 13 40 8.9 69 13 13 Welch 2.093 19
t = 1.228

Sal (0/00) 0 9.0 2.2 3.5 13 0 18.0 9.5 5.3 13 Student’s 2.064 24
t = 4.127

pH 6.2 76 6.7 0.6 12 66 85 7.4 0.6 12 Student’s 2.074 22
t = 2.689

MOA (mg CaCO3/L) 50 775 30.8 21.8 13 4.0 22000.0 17384 60879 13 Welch 2.179 12
t = 1.011

Ca (mg CaCO,/L) 6.2 2460 88.9 80.8 10 23.7 6133 3016 2366 10 Welch 2.201 11
t = 2.690

TFR (mg/L) 100.0 8100.0 35539 27498 13 433.0 21866.0 12377. 7820.2 13 Welch 2.145 14
t = 2.838

Hds (mg CaCO /L) 31.0 8200.0 17360 2969.7 8 92 134333 31240 42360 8 Student’s 2.145 14
t = 0.759

SOC (%) 116 2906 24.09 606 12 651 1490 9.85 2.17 12 Welch 2.160 13
t = 7.667

complex crossed-lamella three and reticulate microstructure.
Microstructure of the inner shell layer (Fig. 5) is always of the
reticulate type.

Exposed tips of secondary lamellae in complex
crossed-lamella one are variably shaped with broad surfaces,
and are oriented almost parallel to the shell surface (Fig. 6).
Exposed tips of secondary lamellae in complex crossed-
lamella two are narrow and also variably shaped, oriented ir-
regularly or obliquely to the shell surface (Fig. 7). Exposed
tips of secondary lamellae in complex crossed-lamella three
are also irregularly shaped, oriented almost perpendicular to
the shell surface (Fig. 8) with lamellae extending farther out
to the inner surface of shell than the neighboring lamellae.
Reticulate shell microstructure consists of loosely to densely
packed thin or thick meshwork that can be granulated (Fig. 9).

The microstructure of the inner shell surface in Area
G can be grouped into irregular blocks (Figs. 10, 11). There
is usually no detectable microstructure that represents a tran-
sition at the presumed ‘‘transition zone’’ (Areas D to F, just
dorsal and adjacent to the pallial line) since this zone is fre-
quently eroded. Thus, the dorsal boundary of the outer shell
layer can be recognized as an elevated border or ridge along
the curved antero-posterior axis. A convenient boundary be-
tween outer crossed-lamellar and inner complex crossed-

Table 2. Lengths of wild and caged Polymesoda caroliniana from
Ocean Springs, Mississippi, whose internal shell surface microstruc-
ture was examined by scanning electron microscopy. Length
measurements are in millimeter (min = minimum, max = maximum,
S = submerged, E = emerged).

Date Mean Standard Range Total
deviation min max clams
examined
(n)
WILD
June 1985 276 8.2 15.3 41.0 20
Sept 1985 29.3 49 20.0 37.6 29
Dec 1985 275 75 16.4 37.0 10
Mar 1986 33.4 45 245 38.5 10
June 1986 33.4 49 24.1 41.6 10
CAGED (S)
Sept 1985 31.4 4.7 23.1 378 10
Dec 1985 34.1 3.3 273 41.5 30
Mar 1986 32.4 3.6 27.7 37.4 10
June 1986 34.7 5.2 29.2 415 10
CAGED (E)
Sept 1985 30.8 3.7 25.2 36.0 10
Dec 1985 29.5 48 23.4 38.0 10

202 AMER. MALAC. BULL. 6(2) (1988)

Fig. 2. Microstructures of inner surface of shell dorsal to the pallial line are grouped into irregular mounds. Mounds represent first order lamellae
[Horizontal field width (HFW) = 352 nm. Fig. 3. Angular view of shell fracture dorsal to the pallial line. Second order lamellae of first order
lamellae are oriented opposite to each other (HFW = 587 um). Fig. 4. Few mounds are visible on rough surface of inner shell (HFW = 587
um). Fig. 5. Irregular layers on surface of inner shell consists of reticulated microstructure (see Fig. 9) (HFW = 293 um).

lamellar shell layers is therefore an eroded groove in place
of an obvious pallial myostracum. This is evident at low
magnifications (Fig. 12), where an apparent transition zone
is seen only at a low magnification. Unlike the usually in-
distinct pallial myostracum, the adductor myostracum is
distinct (Fig. 13). Both pallial (Fig. 14) and adductor (Fig. 15)
myostraca can be traced sandwiched between the two shell
layers.

Ventral to the myostracum (Areas A, B and C), the
microstructures of the inner shell surface of Polymesoda
caroliniana and Corbicula fluminea (Prezant and Tan Tiu, 1985,
1986; Tan Tiu, 1987) are similar, except that no spiral shell
formations were observed in P caroliniana during colder
seasons. Adjacent and ventral to the myostracum, Area C,
the exposed lath tips are irregularly arranged. Area C is often
covered by an organic matrix that render the underlying struc-
tures indistinct. Laths in Area B, dorsal and adjacent to the
area undertucked by the periostracum, are arranged regularly
to form second order lamella. Direction of the second order

lamellae are opposite to that of the adjacent first order lamella.
Microstructure of the inner shell surface of both Area B and
C are referred to as crossed-lamella two (Table 3). Reticulate
microstructure with loosely arranged strands can be observed
at times on Area B. The predominant microstructure of the
inner shell surface in Area A (undertucked by periostracum)
is crossed-lamella one in all three groups. Exposed tips of
secondary lamellae in crossed-lamella one are irregularly ar-
ranged, neither forming rosette nor pseudospiral pattern.
Microstructure C, a collective term of convenience referring
to pseudospiral (Fig. 16) and rosette microstructure in
Polymesoda caroliniana, is similar to that of microstructure
C in Corbicula fluminea, except that in the former, no com-
plete spiral was observed. In P caroliniana, two arc-shaped
secondary lamellae (Fig. 16) can be joined to one another to
form an approximate circular structure. When overlain by
organic matrix, the identity of each arc can be obscured, thus
appearing as a continuous circular flat band. The tertiary
lamellae, composing the hub of the arc secondary lamellae

TAN TIU: POLYMESODA CAROLINIANA SHELL MICROSTRUCTURE 203

are sometimes not aligned, such that the tertiary lamellar tips
protrude at varying lengths into the central space of the cir-
cular structure. Therefore, the shape of the spaces enclosed
by the secondary lamellae vary depending upon the degree
of curvature of the secondary lamellae.

Seasonal and habitat variations in the microstructure
of the inner shell surface of caged and uncaged Polymesoda
caroliniana are summarized in Table 3. Over the 13 month
period, microstructure C in wild clams was absent in June
of 1985 and 1986, and its frequency of occurrence peaks in
March 1986 (Table 3). The frequencies of occurrence of the
following microstructures were highly correlated (r > critical
values at fo95(2)3 = 0.878) in wild clams: microstructure C
negatively correlated with crossed-lamella one and complex
crossed-lamella one. Other microstructures of the inner shell
surface did not show distinct seasonal patterns. Microstruc-
ture C was negatively correlated, whereas complex crossed-
lamella one (r = 0.941) was positively correlated significantly
with temperature of surface water at the time of sampling than
the temperature average per season.

During the four seasons over the 12 month period,
microstructure C in submerged clams was absent in
September 1985 and June 1986, but its frequency of occur-
rence also peaks in March 1986 like that of wild clams (Table
3). Frequency of occurrence of crossed-lamella one was
negatively correlated with that of microstructure C. Other
microstructures of the inner shell surface did not show distinct

Table 3. Temporal and spatial variation of internal shell surface
microstructure in Polymesoda caroliniana, Jackson County, Mississip-
pi. Headings stand for areas of shell examined (first row) and shell
microstructural type (Second row). Shell microstructure abbreviations
are: C, microstructure C; CL, crossed-lamella; CCL, complex crossed-
lamella; Ret, reticulate. Frequency of occurrence expressed in per-
cent, where 0 = 0%, 1 = 1 to 20%, 2 = 21 to 40%, 3 = 41 to 60%,
4 = 61 to 80%, 5 = 81 to 100%.

Area A Areas B-C Areas G-l
Cc CL1 CL2 Ret CCL1 CCL2 CCL3 Ret

Rod and Reel Fishing Camp (wild)

June 1985 0 5 5 0 1 0 5 0
Sept 1985 1 5 5 1 1 3 2 1
Dec 1985 2 3 5 0 0 3 3 1
Mar 1986 3 3 5 1 0 2 1 3
June 1986 0 5 5 0 1 1 1 3
Submerged Area (caged)

Sept 1985 0 5 5 0 2 3 1 1
Dec 1985 1 4 5 1 1 2 2 2
Mar 1986 3 3 4 1 1 2 2 3
June 1986 0 i) 5 0 1 1 1 4

Exposed Area (caged)
Sept 1985 1 5 5 0 0 2 2 2
Dec 1985 3 3 4 1 0 3 0 3

Fig. 6. Complex crossed-lamella one (HFW = 22 um). Fig. 7. Complex crossed-lamella two (HFW = 22 um). Fig. 8. Complex crossed-lamella
three (HFW = 22 um). Fig. 9. Reticulate microstructure (HFW = 22 um).

204 AMER. MALAC. BULL. 6(2) (1988)

Fig. 10. Irregular blocks with smooth surfaces (HFW = 79 um). Fig. 11. Irregular blocks with component secondary lamellae (HFW = 158
pm). Fig. 12. The groove is a convenient boundary between crossed-lamella (above) and complex crossed-lamella (below) (HFW = 790 um).
Fig. 13. Adductor myostracum consists of tall prisms. Remnant of adductor muscle at top of photo (HFW = 31 um). Fig. 14. Organic compart-
ments of pallial myostracum are sandwiched between naturally eroded shell layers (HFW = 16 um). Fig. 15. Adductor myostracum is sand-

wiched between outer shell layer, crossed-lamella (above), and inner shell layer, complex crossed-lamella (below) (HFW = 32 um).

seasonal pattern.

In exposed clams, available data on microstructure of
the inner shell surface for September and December 1985
indicated that increase in the frequency of occurrence of
microstructure C and decrease in crossed-lamella one were
similar to those in wild clams. However, complex-crossed
lamella one that was consistently present in submerged
clams, present only in June and September in wild clams,
were absent in exposed clams.

CONDITION INDEX AND ORGANIC CONTENT
OF SHELL

Average percentages (+ one standard deviation, n) of
condition indices in wild (June 1985 = 3.30 + 0.97,n = 44,
Sept 1985 = 2.64 + 0.95,n = 45, Dec 1985 = 2.87 + 0.96,
n = 49, Mar 1986 = 4.00 + 1.17, n = 50, June 1986 = 4.38
+ 0.99, n = 50), and submerged bivalves (Sept 1985 = 3.64
+ 1.53, n = 37, Dec 1985 = 4.60 + 1.11,n = 11, Mar 1986
= 6.22 + 1.06,n = 4) varied significantly as tested by ANOVA
(Tables 4). In exposed clams, samples were available only for
September 1985 (5.14 + 0.80, n = 6). The Tukey test indicates
that the condition index in wild clams can be divided into three
groups; June-1985, September 1985, and March 1986-June
1986. Tukey test could not determine how the December con-
dition index was related to either June or September 1985
groups (at least one type II error has been committed) (Table
4). Condition indices in submerged clams can be divided in-
to September 1985 and March 1986 groups. A Tukey test could
not determine how the December condition index was related

ers

ao, ne P44; Wala, 4 SIN Son
thy — ay Sa be

%,

Fig. 16. Arc-shaped secondary lamellae can join to form a hub in
Area A (area undertucked by the periostracum) during cooler months

(Dec. and Mar.) (HFW = 16 um).

to either September or March groups (Table 4).

The condition index in December 1985 was significant-
ly different among wild, submerged and exposed clams.
Moreover, the Tukey test indicated that the condition index in
wild clams was different from submerged and exposed clams
(Table 5). A pairwise comparison of average condition index
in wild and submerged clams also indicated that condition
index in September (Welch t = 3.522 > toos,2)7 = 2.365) and
March (Student’s t = 3.661 > toos;2)52 = 2.007) were
significant.

Analysis of variance of average percentages (+ one
standard deviation, n) of organic content of shell did not show

TAN TIU: POLYMESODA CAROLINIANA SHELL MICROSTRUCTURE 205

Table 4. Temporal variation of means (x) of condition index (Cl) and shell organic content (SOC) in Polymesoda caroliniana from Rod and
Reel Fishing Camp (wild) and submerged area (submerged), Ocean Springs, Jackson County, Mississippi.

Analysis of Variance (one-way) Tukey test

Computed value Table value
6/85 9/85 12/85 3/86 6/86 N D F F N D Overall conclusion
Cl
Wild 330 2.64 287 400 4.38 4 235 25.83" 2.85 4 200 X1 # Xo # Xq4 = Xs5
Submerged 364 460 6.22 — 2 49 6.95" 4.01 2 45 X1 # X3
SOC
Wild 2.72 2.83 2.87 2.84 2.75 4 236 1.19 2.85 4 200 not necessary
Submerged 2.74 258 2.73 — 2 47 1.45 4.01 2 47 not necessary

*The ratio of the group mean square over the error mean square (F) with N and D degrees of freedom respectively is significant at a = 0.05.
When degrees of freedom fall between two table values, the lower value is used. Cl and SOC are in %. Subscripts for x correspond to the
order (left to right) of the means. Absence of available data is represented by blank spaces (before) and — (during) the sampling period.

Table 5. Tukey test among the average condition indices (Cl) of Polymesoda caroliniana
in three different habitats for December 1985. Computed studentized range (q) = (Xp
- Xa) + standard error. Critical value = 3.399 at a = 0.05, degrees of freedom = 65
= 60, and total number of means tested = 3(S = submerged, E = exposed area).

Habitat Wild Caged (S) Caged (E)
Samples ranked by means (i) 3 2 1
Ranked sample means (x;) 2.87 4.60 5.14
Comparison Difference Standard

(B vs A) (Xp - Xa) error q Conclusion

1vs 3 2.27 0.26 8.73 Reject Ho: x; = x3

1 vs 2 0.54 0.23 2.35 Accept Ho: x; = X2

2vs 3 1.73 0.32 5.41 Reject Ho: x2 = x3
Overall conclusion: xX; = Xo # X3

significant seasonal differences in wild (June 1985 = 2.72
+ 0.41,n = 48, Sept 1985 = 2.83 + 038, n = 45, Dec 1985
= 2.87 + 056,n = 49, Mar 1986 = 2.84 + 0.40,n = 50,
June 1986 = 2.75 + 0.23,n = 49) (Table 4) and submerged
clams (Sept 1985 = 2.74 + 0.29, n = 35, Dec 1985 = 2.58
+ 0.16, n = 11, Mar 1986 = 2.73 + 0.13, n = 4) (Table 4).
In exposed clams, samples were available only for Sept 1985
(2.70 + 0.19, n = 8). Average percentages of organic con-
tent of shell in Dec for wild, submerged and exposed clams
were not significantly different as indicated by ANOVA (F =
174 < Fo.05(2)2,65 = Fo.05(2)2,60 = 3.93). Moreover, pairwise
comparison of shell organic content in Sept (Student’s t =
1.167 < to.05(2)78 = 1.991) and Mar (Welch t = 1.241 <
1o05(2)9 = 2-262) between wild and submerged clams was
not significant.

DISCUSSION

Among the microstructures of the inner shell surface
in Table 3, complex-crossed lamella one reflects habitat dif-
ferences. Complex-crossed lamella one was present through-
out the year in submerged clams, present only during June
and September in wild clams, and absent in exposed clams.
Environmental conditions in these three habitats were dif-
ferent. The submerged area was less stressful than the ex-
posed area. Of 360 individuals in each group (exposed and
submerged), 45% were recovered alive from the submerged

while only 13% were recovered from the exposed area.
“Stress can be said to occur when physiological (or other)
processes are altered in such a way as to render the individual
less fit for survival’ (Bayne, 1980). Moreover, shell formation
is costly. Shell formation involves ion transport, protein syn-
thesis and sequences of physiological processes (Wilbur and
Saleuddin, 1983). ‘‘Healthier’’ clams would therefore be ex-
pected to have more energy allocated for shell formation and
maintenance than less ‘‘healthy’”’ clams.

Among the internal shell surface microstructures
observed in this study, microstructure C and crossed-lamella
one were the most conservative in the sense that seasonal
patterns (frequency of occurrence) among the three groups
wild, exposed and submerged clams were almost similar
despite differences in habitat. Frequency of occurrence of
microstructure C is inversely associated with temperature of
water surface at time of sampling in wild clams, and with
average temperature of water surface in submerged clams.
Difference in time response could be due to temperature
stability provided by water to submerged clams in a continually
submerged habitat.

Other than what has been discussed above, the
seasonal and habitat variation nor the factor associated with
the presence and frequency of occurrence of microstructure
in the inner shell surface is not clear. Reticulate microstructure
did not show seasonal variation instead increased in all three
types throughout the experimental period. This microstruc-

206 AMER. MALAC. BULL. 6(2) (1988)

ture is possibly a common response to altered environment
induced by several factors.

Palmer (1983) reported that production of skeletal
organic matrix can be more ‘‘demanding metabolically than
the crystalization of calcium carbonate.’ Therefore, high
amount of organic content of shell is expected to occur dur-
ing the time when clams are ‘‘healthiest’’. However, organic
content of shell did not show significant seasonal variation.
Possibly the difference if any during the study period were
diluted by the total content through the life of the animal as
suggested by an anonymous reviewer of this paper.

In view of the data presented here and elsewhere (Tan
Tiu, 1987; Prezant and Tan Tiu, 1986), it seems that micro-
structure of the inner shell surface outside the pallial line,
especially on Area A (area undertucked by periostracum),
although showing seasonal variation and slight habitat varia-
tion, is characteristic of some species. That is, while Corbicula
fluminea can form spiral shell microstructures, Polymesoda
caroliniana cannot. Shell outside the pallial line could indeed
be a conservative characteristic of the species, and therefore
could be used in taxonomic or phylogenetic analyses. On the
other hand, shell microstructure beyond basic components
inside the pallial line, by virtue of its greater variability
(changes in shell ultrastructure due to formation, modifica-
tion, dissolution, etc.) as a reflection of changes in shell
physiology due to environmental changes, can be used for
taxonomic purposes only if ontogeny and environmental
history are known. The variability of shell ultrastructure out-
side the pallial line in other corbiculids needs further study.

ACKNOWLEDGMENTS

| am grateful to Drs. Robert S. Prezant, Clement L. Counts,
lll, J. Gaylord Carter, Mrs. Rebecca Bogart and two anonymous
reviewers for the review of this manuscript, and to Mr. Noel D’mello,
Dr. Richard Heard, Mr. Thomas Rogge, Miss Alene Minchew, Dr.
Kumar Prasanna, Dr. Robert S. Prezant and Mr. Sheau-Yu Shu for
help in sample collection; Dr. Raymond Scheetz for help with scan-
ning electron microscopy; Mr. Tom Smoyer for printing figure 13; Ms.
Jackie Van Mort of the Rod and Reel Fishing Camp for allowing me
to collect samples on their property; Dr. Harold D. Howse for allow-
ing me to transplant clams into a marsh on the Gulf Coast Research
Laboratory property. This study was supported by a National Capital
Shell Club Scholarship, a grant from Sigma Xi and aid from the
Department of Biological Sciences, University of Southern Mississippi,
U.S.A., and Research and Faculty Development from the University
of San Carlos, Philippines. Publication cost supported by Harbor
Branch Oceanographic Institution, Inc. (Contribution Number 623).

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Lutz, R. A. and D. C. Rhoads. 1980. Growth patterns within the
molluscan shell. /n: Skeletal Growth of Aquatic Organisms.
Rhoads, D. C. and R. A. Lutz, eds. pp. 203-254. Plenum Press,
New York.

Palmer, A. R. 1983. Relative cost of producing skeletal organic matrix
versus Calcification: evidence from marine gastropds. Marine
Biology 75:287-292.

Prezant, R. S. and A. Tan Tiu. 1985. Comparative shell microstruc-
ture of North American Corbicula (Bivalvia: Sphaeriacea).
Veliger 27(3):312-319.

Prezant, R. S. and A. Tan Tiu. 1986. Spiral crossed-lamellar shell
growth in the bivalvia Corbicula fluminea (Mollusca: Bivalvia).
Transactions of the American Microscopical Society
105(4):338-347.

Prezant, R. S. and K. Chalermwat. 1983. Environmentally induced
changes in shell microstructure of the Asiatic clam Corbicula.
American Zoologist 23(4):914.

Rhoads, D. C. and G. Panella. 1970. The use of molluscan shell growth
patterns in ecology and paleoecology. Lethaia 3:143-161.

Tan Tiu, A. 1987. Influence of environment on shell microstructure
of Corbicula fluminea and Polymesoda caroliniana. Doctoral
Dissertation, University of Southern Mississippi, Hattiesburg.
148 pp.

Tan Tiu, A. and R. S. Prezant. 1987. Shell microstructural responses
of Geukensia demissa granosissima (Mollusca: Bivalvia) to con-
tinual submergence. American Malacological Bulletin
5(2)173-176.

Taylor, J. D., W. J. Kennedy and A. Hall. 1969. The shell structure
and mineralogy of the Bivalvia. Introduction, Nuculacea-
Trigonacea. Bulletin of the British Museum (Natural History)
Zoology 22(9):255-294.

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mineralogy of the Bivalvia. II. Lucinacea-Clavagellacea, Con-
clusions. Bulletin of the British Museum (Natural History)
Zoology 22:235-294.

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vertebrate. Progress in Crystal Growth Characterization 4:99-147.

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Mollusca. Vol. 4. Physiology, part 1. Wilbur, K. M., ed. pp.
235-287. Academic Press, Inc., New York.

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718 pp.

Date of manuscript acceptance: 13 January 1988.

THE USE OF ARM SUCKER NUMBER IN
OCTOPODID SYSTEMATICS (CEPHALOPODA: OCTOPODA)

RONALD B. TOLL
DEPARTMENT OF BIOLOGY
THE UNIVERSITY OF THE SOUTH
SEWANEE, TENNESSEE 37375, U. S. A.

ABSTRACT

The average total number of suckers per arm for twelve species of octopodine cephalopods
is presented in terms of the rate of sucker addition during growth. These data are shown to be useful
for systematic analysis. The rate of sucker addition displays positive allometry relative to arm growth
in early stages of development. Sucker addition slows to become negatively allometric in subadults
and adults. New sucker morphogenesis ceases in the late stages of growth in some taxa resulting
in an apparent species-specific sucker number. The hectocotylized arm displays a similar ontogenetic

pattern of sucker addition.

Based on presumed reproductive isolation, general robustness, average arm sucker count (AASC),
hectocotylized arm sucker count (HASC), and brooding mode, Scaeurgus patagiatus Berry, 1913 is
removed from the synonomy of S. unicirrhus Orbigny, 1840 and is considered to be a separate species.

The total number of suckers on the arms of octopods
is perhaps the second most saliant meristic feature after the
nominal character of the order (Octopoda eight legs),
which, being invariant among normal specimens, is of no
systematic value among subordinal taxa. Counts of arm
suckers occasionally were included as part of systematic
descriptions and biological investigations of octopodid taxa,
e.g. Férussac and Orbigny (1834-48), Troschel (1857), Verrill
(1882), Jatta (1896), Naef (1923), Winckworth (1928), Sasaki
(1929), and Boletzky (1975), but most contemporary workers
have ignored this character. Furthermore, | am unaware of
any published account that compares octopodids, either inter-
or intraspecifically, based on sucker counts or that employs
these data in broader comparative studies at any taxonomic
level. The limited use of either total number of arm suckers
or the number of sucker rows in systematic treatments of oc-
topod taxa is difficult to comprehend. The situation can at best
be rationalized by appreciating the time required to count the
suckers on each arm of the numerous specimens required
to construct a significant data base. The total number of
suckers per individual can range from several hundred to
several thousand depending on species and maturity.

Most recently, Roper and Voss (1983), in their prece-
dent setting guidelines for the description of cephalopod taxa,
included arm sucker count (ASC) as a minimal requirement
for the adequate taxonomic description of octopodids.
However, not one of the three papers cited by these authors

as exemplary in octopod systematics include ASC data.

The arm sucker count of hatchling octopuses has been
used as a specific-level systematic character (see Boletzky,
1977, 1984; Hochberg et a/., in press). The potential systematic
value of arm sucker count in post-hatchling to adult octopodids
remains inadequately investigated, an attribute shared by a
large suite of other meristic and morphometric characters (e.g.
gill lamellae number, penis morphology, alimentary tract
anatomy, stellate ganglion morphology, etc.). The evaluation
of ontogenetic rates of sucker morphogenesis also has been
largely ignored, except for the most basic premises that lar-
val octopods, whether benthic or planktonic, hatch with
relatively few suckers compared to adults and that this sucker
number increases with growth.

This paper presents a preliminary systematic survey
of arm sucker counts in octopodine cephalopods. These
results strongly suggest that average arm sucker counts can
be valuable in systematic studies of the Octopodinae. The
results also indicate that the rate of addition of new suckers
shows a decidedly positive allometry with respect to arm
growth in small animals. As a result, octopodids of relatively
small body size precociously attain the majority of the adult
complement of suckers. This phase is followed in late juveniles
or early adults by negative allometry with a near to total cessa-
tion of addition of new suckers during the later stages of arm
growth.

American Malacological Bulletin, Vol. 6(2) (1988):207-211

207

208 AMER. MALAC. BULL. 6(2) (1988)

MATERIALS AND METHODS

The procedure used to count suckers was as follows.
Suckers on each arm were counted starting at the mouth and
moving to the tips. The relatively large suckers on the prox-
imal two-thirds to three-quarters of the arm were counted us-
ing the unaided eye or an illuminated magnifier and passing
a needle probe down the arm. Distally, where the suckers can
be minute and densely packed, suckers were counted using
a binocular microscope. A fine dissecting or insect pin was
inserted into the arm as a marker during the transition be-
tween the two counting procedures. In all cases, sucker
rudiments (anlagen), which appear as minute dome-like pro-
jections at the distal extremities of the arms, were counted
as complete suckers. Suckers that were obviously missing,
lost in combat with predators or prey or during capture, were
counted as if present. Only complete arms with the entire

Ls)
oO

Oo
oO

AASC or HASC
oD @
oS

L
oO

AASC or HASC

AASC or HASC

AAL or HAL (mm)

Figs. 1-6. Scattergrams of AASC vs. AAL and HASC vs. HAL for six species of Octopodinae [O = unmodified (nonhectocotylized) arms;

distal tip intact were used to obtain sucker counts. Complete
arms were considered ‘available’ and this term is used below.
Arms regenerating from injury were excluded from considera-
tion. To expedite counting, the number of sucker rows were
counted and this value doubled to obtain the total sucker count
for each individual arm. In cases of irregular sucker place-
ment, acommon artifact of preservation, this procedure could
not be employed and it was necessary to count each sucker
individually.

Arm lengths were measured using traditional methods
with mechanical dividers and standard millimeter rules, follow-
ing the guidelines re-established by Roper and Voss (1983).
Average arm length (AAL) is the mean length of all available
arms, with the exception of the hectocotylus. Average arm
sucker count (AASC) is the mean of the number of suckers
of all available arms with the exception of the hectocotylus.
Both values are expressed to the nearest integer. Values

fe) 50 100 150 200 250 300 350 400 450 500 550. 600

AAL or HAL (mm)

= hectocotylized arm; each symbol represents a single animal]. Fig. 1. Octopus burryi. Fig. 2. Octopus hummelincki. Fig. 3. Octopus selene.
Fig. 4. Octopus digueti. Fig. 5. Octopus defilippi. Fig. 6. Octopus dofleini.

TOLL: OCTOPODID SUCKER NUMBER 209

reported here are from individual animals with at least two
available arms. Hectocotylus arm length (HAL) and hec-
tocotylized arm sucker count (HASC) were separately record-
ed. All specimens examined were preserved in alcohol and
most, if not all, were previously fixed in formaldehyde.
Shrinkage of the arms is assumed to have occurred as a result
of this chemical treatment (see Andriguetto and Haimovici,
1988). Scattergrams and statistical regression analyses were
performed using a MacIntosh Plus© micro-computer with the
statistical program Statworks 512+ ©.

RESULTS AND DISCUSSION

AAL, AASC, HAL, and HASC data from twelve species
of octopodines are plotted in figures 1-11: Octopus burryi Voss;
O. hummelincki Adam (=O. filosus Howell); O. selene Voss; O.
digueti Perrier and Rochebrune; O. defilippi Verany; O. dofleini

(Wulker); Pteroctopus tetracirrhus (delle Chiaje); Robsonella
fontanianus (Orbigny); Scaeurgus unicirrhus Orbigny; S.
patagiatus Berry; Hapalochlaena cf. maculosa (Hoyle); Cisto-
pus indicus (Orbigny). Second-order regression lines are in-
cluded for all data sets where n =5 (except A. fontanianus).

Preliminary regression analyses used each available
arm on all animals as a separate datum, with the exception
of the hectocotylus. The resultant scattergrams, combined
with a basic understanding of octopod growth, showed that,
for any one animal, arm sucker counts and arm lengths are
autocorrelated, thereby jeopardizing the statistical validity of
the regression. Individual averaging of the two data sets from
each animal greatly reduced the size of the resulting data sets
but served to enhance their robustness.

Larval and small juvenile specimens are absent from
the present analyses, a reflection of the relative lack of
representation of small individuals in museum collections and

oO
1%)
<
x
ro)
oO
i?)
¢
<
oO
a
<
=
ro)
oO
Yn
<
<
200
180
160
B 140 O
= 120 o
100
oO
80
B 6
Zz 0)
< 40
i 11
ie)
ie) 5:0) 100 150 200 250 300 350 400
AAL or HAL (mm)
Figs. 7-11. Scattergrams of AASC vs. AAL and HASC vs. HAL for six species of Octopodinae [O = unmodified (nonhectocotylized) arms;

L) = hectocotylized arm; each symbol represents a single animal]. Fig. 7. Pteroctopus tetracirrhus. Fig. 8. Robsonella fontanianus. Fig. 9.
Scaeurgus unicirrhus (darkened symbols), Scaeurgus patagiatus (open symbols). Fig. 10. Hapalochlaena cf. maculosa. Fig. 11. Cistopus indicus.

210 AMER. MALAC. BULL. 6(2) (1988)

the difficulty of identification of young octopodines. Therefore,
the size ranges of some taxa included here are restricted to
sub-adults and adults. Nonetheless, compared to the rate of
arm growth (as a linear measurement), addition of arm
suckers shows a distinct positive allometry during early growth
stages. Small, presumably young, individuals have a
disproportionately large percentage of their full adult comple-
ment of suckers. In Octopus burryi (Fig. 1), animals from 44-59
mm AAL had attained an average of 80.6% of the mean sucker
count of animals from 98-119 mm AAL. Similar trends are seen
in O. hummelincki (Fig. 2), O. diqgueti (Fig. 4), O. defilippi (Fig.
5), Scaeurgus patagiatus, S. unicirrhus (Fig. 9), Hapalochlaena
cf. maculosa (Fig. 10) and Cistopus indicus (Fig. 11). AASC
in Octopus selene (Fig. 3), Pteroctopus tetracirrhus (Fig. 7),
and Robsonella fontanianus (Fig. 8) was statistically invariant
over the size ranges reported here (F = 2.29, 0.71, and 1.81,
respectively; p >.05). Most of arm suckers in small individuals
are rudimentary or minute and densely packed along the arm
tip. Arm growth proceeds by elongation and expansion at the
tips, while the anlagen located there enlarge and become
more widely spaced. Data from larger specimens show a
negative allometric relationship between sucker addition and
arm growth. Indeed, in the final stages of arm growth, very
few if any new sucker anlagen are added and the number
of sucker rudiments and minute suckers is reduced as the
suckers enlarge to reach their definitive sizes.

Average sucker number appears to reach a maximum
value in each species in an apparent display of determinant
growth. These maxima differ among the species examined;
however, while they are presumed to be genetically deter-
mined, it seems unlikely that future study will elucidate non-
overlapping species-specific values because of the large
number of octopodine taxa. Average sucker number data can,
however, assist in identification and taxonomic delineation of
taxa from restricted geographical areas or that are otherwise
morphologically similar (see below). Furthermore, the reduc-
tion of the number and density of rudimentary suckers along
the distal tip of the arms could be valuable in recognizing en-
vironmentally induced precocious onset of sexual maturation
in undersized individuals, a matter of considerable importance
in studies of the structure of wild populations as well as ar-
tificially induced maturation of laboratory cultured animals.
The change from positive to negative allometry could coin-
cide with important ecological or developmental changes yet
to be recognized.

It is well known that among octopodid taxa the char-
acteristic length of the arms varies with respect to body size
(mantle length). Data presented here suggest that AASC and
HASC also vary with respect to arm length among different
taxa, apparently a function of both sucker size and linear den-
sity (Compare Figs. 5, 6, 8).

The hectocotylized arm of males presents a special case.
Without exception HASC was lower than AASC for all in-
dividuals of all taxa examined. The rudimentary calamus and
ligula form early in ontogenetic development from the distal
tip of the arm which is partially devoid of sucker anlagen. By
the onset of calamus and ligula morphogenesis, sucker mor-
phogenesis has slowed considerably and soon ceases. There-

fore, the total sucker complement of the hectocotylized arm
is less than, and is reached earlier in ontogeny than, any of
the nonhectocotylized arms. Also, the most distal suckers are
larger than those of the nonhectocotylized arms. HASC could,
therefore, be a better taxonomic character despite its restric-
tion to male individuals.

Each species appears to be characterized by a nar-
row range of values for HASC but, as with AASC, the large
number of octopodid species probably precludes unique
species-specific values. As with AASC, HASC also could be
significant in restricted taxonomic applications (see below).

Reduction in length of the hectocotylized arm in com-
parison to the fellow arm of some taxa is well documented
among the octopods. Also, the length of the modified portion
of the arm varies among species, ranging from about 1 to 25%
of the arm length. It is expected therefore, that the HASC
varies among taxa independently of either general body size
or lengths of the nonhectocotylized arms.

Analysis of AASC and HASC from a large collection
of Scaeurgus spp. (n=44) (Fig. 9) provided unexpected and
taxonomically provocative results. The Atlantic Ocean (Florida,
Caribbean, Mediterranean) and Pacific Ocean (Hawaii, Japan)
populations show distinctly different and non-overlapping
values of AASC for same-sized individuals and of HASC for
all-sized individuals. Scaeurgus unicirrhus was originally
described by d’Orbigny (1840) from the Mediterranean Sea.
Berry (1913) erected Scaeurgus patagiatus from the Hawaiian
Islands, supplementing his description the following year
(Berry, 1914). Berry recognized the slightly larger size of the
Hawaiian form and the zoogeographic (reproductive) separa-
tion of the two populations. He felt this was sufficient grounds
to separate them at the species level. Robson (1929) synon-
omized S. patagiatus with S. unicirrhus, remarking that all dif-
ferences between the two were insignificant except for the
greater arm lengths in the Pacific form. Subsequently S. uni-
cirrhus has been reported from the Indian Ocean (Robson,
1929) and the Western Atlantic Ocean (Voss, 1951). It has not
been reported from the eastern Pacific Ocean. The genus is
restricted to tropical and warm temperate waters.

The Atlantic and Pacific forms of Scaeurgus are and
probably have been reproductively isolated for an extended
period of time, at least since the last closure of the Isthmus
of Panama. The two populations differ substantially in max-
imum size, arm robustness, AASC, and HASC. Furthermore,
the Pacific form is reported to brood its eggs by holding them
within the web (W. Van Heukelem, pers. comm.; also see
Boletzky, 1984), apparently an ususual behavior among oc-
topodines (see Wells, 1978; Mangold, 1987). The more com-
mon practice of cementing the eggs to the substratum is
displayed by the Atlantic form (Boletzky, 1984). | believe that
the Pacific form merits the specific delineation recognized by
Berry and correctly should be called Scaeurgus patagiatus
Berry, 1913.

The relative simplicity of counting arm suckers facili-
tates routine examination even by inexperienced workers.
Replicate sucker counts performed by novice assistants in
the present study were routinely close, typically with errors
of 2% or less. The greatest source of potential error involves

TOLL: OCTOPODID SUCKER NUMBER 211

the sucker rudiments on the arm tips. Some experience helps
to standardize the counting procedure to include all true
rudiments while excluding artifactual convolutions of the oral
surface of the arm caused by fixation and/or preservation.

The use of total number of arm suckers rather than the
number of sucker rows could be seen as arbitary in view of
the biserial sucker arrangement found in all octopodines. In-
deed, in many cases sucker rows were counted and multiplied
by two to obtain total sucker number. However, the uniformi-
ty of the biserial arrangement often is lost in portions of some
arms in many individuals. Also, in larval specimens and in
adults of some taxa, the first several adoral suckers are
uniserial (Howell, 1868; Naef, 1923). Finally, the use of total
counts will facilitate future comparisons with octopodids with
uniserial sucker arrangements (e.g. Eledoninae), and does
not suggest an unwarranted homology between a single row
of suckers of the biserial octopodines and individual suckers
of the eledonids and related groups.

ACKNOWLEDGMENTS

The following persons and institutions kindly provided oc-
topodine material studied here: Clyde F. E. Roper and Michael J.
Sweeney, National Museum of Natural History, Smithsonian Institu-
tion; Gilbert L. Voss, Rosenstiel School of Marine and Atmospheric
Science; Brian Hartwick, Simon Fraser University; Wolfgang Zeidler,
South Australian Museum; Janet Voight, University of Arizona. Their
assistance is deeply appreciated.

This study was supported by a grant from the National Science
Foundation (BSR-85084339), a Postdoctoral appointment to the Divi-
sion of Mollusks, National Museum of National History, Smithsonian
Institution by the Smithsonian Office of Fellowships and Grants, and
a faculty research grant and general support from The University of
the South. This support is gratefully acknowledged.

Discussions with Gilbert Voss and Janet Voight were helpful
to the writer. Constructive criticisms offered by Clyde Roper, Michael
Vecchione, and Janet Voight upon earlier drafts of this paper were
helpful. Sigurd von Boletzky provided additional information and con-
structive criticisms upon a later version. This manuscript was typed
by the staff of the Word Processing Center at The University of the
South.

LITERATURE CITED

Andriguetto, J. M. and M. Haimovici. 1988. Effects of fixation and
preservation methods on the morphology of a loliginid squid
(Cephalopoda: Myopsida). American Malacological Bulletin
6(2):213-217.

Berry, S. S. 1913. Some new Hawaiian cephalopods. Proceedings of
the United States National Museum 45:563-566.

Berry, S. S. 1914. The Cephalopoda of the Hawaiian Islands. Bulletin
of the United States Bureau of Fisheries 32:257-362.
Boletzky, S. von. 1975. Le développement d’Eledone moschata
(Mollusca, Cephalopoda) élevée au laboratoire. Bulletin de la

Société Zoologique de France 100(3):361-367.

Boletzky, S. von. 1977. Post-hatching behaviour and mode of life in
cephalopods. Symposia of the Zoological Society of London
No. 38:557-567.

Boletzky, S. von. 1984. The embryonic development of the octopus
Scaeurgus unicirrhus (Mollusca, Cephalopoda). Additional data
and discussion. Vie et Milieu 34(2/3):87-93.

Férussac, M. and A. d’Orbigny. 1834-1848. Histoire Naturelle Generale
et Particuliére des Céphalopodes Acétabuliferes Vivant et
Fossiles. Paris. 366 pp.

Hochberg, F. G., M. Nixon, R. Toll and R. Young. (In press). The Order
Octopoda Leach, 1818. /n: Larval and Juvenile Cephalopods.
Proceedings of the First International Cephalopod Advisory
Council Workshop, Banyuls, France, June, 1985.

Howell, S. B. 1868. Descriptions of two new species of Cephalopods.
American Journal of Conchology 3:239-241.

Jatta, G. 1896. Cephalopodi. Fauna und Flora des Guifs von Neapel.
R. Friedlander and Sohn, Berlin. 268 pp.

Mangold, K. 1987. Reproduction. /n: Cephalopod Life Cycles, Vol. 11.
Comparative Reviews. P. R. Boyle, ed. pp. 157-200, Academic
Press, London.

Naef, A. 1923. Die Cephalopoden. Fauna et Flora des Golfes de
Neapel 35 (I), 863 pp.

Orbigny, A. d’. 1840. In: Ferussac, M. and A. d’Orbigny. Histoire
Naturelle Générale et Particuliére des Céphalopodes
Acétabuliferes Vivant et Fossiles. Paris. 366 pp.

Robson, G. C. 1929. A Monograph of the Recent Cephalopoda, Part
|, Octopodinae. British Museum, London. 236 pp.

Roper, C. F. E. and G. L. Voss. 1983. Guidelines for taxonomic descrip-
tions of cephalopod species. Memoirs of the National Museum
of Victoria No. 44:49-63.

Sasaki, M. 1929. Monograph of the dibranchiate cephalopods of the
Japanese and adjacent waters. Journal of the College of
Agriculture, Hokkaido University 20 (suppl. no.), 357 pp.

Troschel, H. 1857. Bemerkungen uber die Cephalopoden von
Messina. Archiv Naturgesch, Berlin 23:41-76.

Verrill, A. E. 1882. The Cephalopods of the Northeastern Coast of
America. Report of the United States Fishery Commission for
1879:211-455.

Voss, G. L. 1951. A first record of the cephalopod, Scaeurgus uni-
cirrhus, from the Western Atlantic. Bulletin of Marine Science
of the Gulf and Caribbean 1 (1):64-71.

Wells, M. J. 1978. Octopus-Physiology and Behaviour of an Advanced
Invertebrate. Chapman and Hall, London. 417 pp.

Winckworth, R. 1928. The hectocotylus of Octopus octopodia (L.) Pro-
ceedings of the Malacological Society, London 18:49-50.

Date of manuscript acceptance: 1 April 1988.

RESEARCH NOTE

EFFECTS OF FIXATION AND PRESERVATION METHODS
ON THE MORPHOLOGY OF A LOLIGINID SQUID
(CEPHALOPODA: MYOPSIDA)

JOSE MILTON ANDRIGUETTO JR.
ZOOLOGY DEPARTMENT, UNIVERSIDADE FEDERAL DO PARANA,
CX.P. 3034, CEP 80.001, CURITIBA, PR., BRAZIL

AND

MANUEL HAIMOVICI
OCEANOGRAPHY DEPARTMENT, FUNDACAO UNIVERSIDADE DO
RIO GRANDE, CX.P. 474, CEP 96.200, RIO GRANDE, RS., BRAZIL

ABSTRACT

The effects of freezing, fixation and preservation in 70% ethanol or 10% formalin for periods
up to 46 months on body morphometry of Loligo sanpaulensis Brakoniecki, 1984, were investigated.
Significant morphometric changes were observed, mainly between previously frozen and non-frozen
specimens. Some forms of long-term preservation produced further, statistically significant changes.
Long-term preservation increased variability of individual effects, widening confidence limits of most
indices. Fresh squids or material recently fixed in a standard way should be used for population studies
if there is no previous knowledge of the effects of fixation techniques on specific measurements.

Body proportions frequently are used as criteria for
distinguishing groups of organisms in terms of species or
populations. Morphometric indices are calculated from soft
part measurements that are more subject to changes due to
fixation and preservation in cephalopods, than, for example,
in crustaceans and vertebrates. Therefore, care must be taken
to recognize real differences as distinct from those caused
by processing techniques.

In loliginids with worldwide distributions and closely
related species and subspecies, morphometric indices have
been used to identify and classify groups in taxonomic studies
(Cohen, 1976; Voss, 1977; Juanico, 1979) as well as to
distinguish stocks or subpopulations (Kashiwada and
Recksiek, 1978; Juanico, 1979). Some papers, notably Cohen
(1976), compared short-term effects on squid morphometric
indices of refrigeration, fixation in 10% formalin, and preser-
vation in isopropyl! alcohol. Our paper deals with short and
long-term changes on loliginid squids fixed and preserved in
10% formalin and 70% ethanol, with and without previous
freezing.

MATERIALS AND METHODS

The effects of fixation and preservation on
measurements and consequently on morphometric indices
were analysed on samples of Loligo sanpaulensis Brakoniecki,
1984, collected in a bottom trawl survey off Rio Grande do
Sul, Brazil, in 1983 (see Haimovici and Andriguetto Jr., 1986).
L. brasiliensis Blainville, 1823, was the name most common-
ly applied to the common loliginid in Brazilian waters in the
majority of papers published in South America (eg.
Castellanos, 1967; Juanico, 1980; Figueiras and Sicardi, 1980:
Vigliano, 1985). Brakoniecki (1984) considered L. brasiliensis
a nomen dubium, since the holotype no longer exists and
because the original description was inadequate and could
refer to any of the species of Loliginidae of the Southwest
Atlantic (see also Voss, 1974).

Eighty-six specimens were measured within two hours
of capture. Then, 40 specimens were frozen, 20 fixed in 10%
buffered formalin in sea water and 26 fixed in 70% ethanol.
The frozen individuals were thawed and measured again

American Malacological Bulletin, Vol. 6(2) (1988):213-217

213

214 AMER. MALAC. BULL. 6(2) (1988)

Table 1. Methods and length of treatments of Loligo sanpaulensis.

Treatment Number of Mantle length Length of treatment
animals range (mm) Short-term Long-term
(days) (months)
10% Formalin 20 36-104 55 46
70% Ethanol 26 38-112 50 46
Freezing 40 52-88 40 46
Freezing; then 10% formalin 20 52-79 40-40 46
Freezing; then 70% ethanol 20 60-88 40-40 46

after 40 days. Half were transferred to 10% formalin and half
to 70% ethanol. All measurements were repeated after 46
months of preservation in the corresponding fixatives (Table
1). These procedures can be considered fixation and preser-
vation techniques as defined by Roper and Sweeney (1983).

Measurements taken were: 1, mantle length (ML); 2,
fin length (FL), from posterior mantle tip diagonally to the in-
sertion of anterior left border; 3, fin width (FW); 4, arm length
(AL), length of third left arm, measured from its tip to the
anterior margin of left eye; 5, length of extended left tentacle
(TL), measured as for the arm; 6, eye diameter (ED).
Measurements 1, 3 and 6 follow Roper and Voss (1983). The
corresponding indices were calculated as percentages of the
mantle length, e.g. TLI, ALI, FLI, FWI, EDI (Roper and Voss,
1983).

COMPARISONS BEFORE AND AFTER
TREATMENTS

Ratios of measurements and indices before and after
fixation, and after almost 4 years of preservation are shown
in figure 1. Values of 1.0 indicate no change, higher values
indicate distension and lower values indicate contraction.
Ninety-five-percent confidence intervals were calculated and
Student’s ‘‘t’’ tests were performed to show significant dif-
ferences between rate values and the value of one.

Fixation in formalin increased FW, FWI and FLI, and
reduced TL and TLI. Formalin preservation reduced FL, TL
and TLI. Fixation and preservation in ethanol reduced all
measurements and indices, except FL and FLI for fixation,
and AL, ALI and FLI for preservation.

Freezing reduced mantle and fin length, and increased
the other measurements. Posterior fixation in ethanol or for-
malin reduced significantly all measurements. The only in-
dex reduced was FWI, after freezing/formalin fixation. Fur-
ther changes occurred for all indices after long-term
preservation.

COMPARISONS AMONG TREATMENTS

Growth of Loligo sanpaulensis within mantle length
ranges of our samples was shown to be allometric by Vigliano
(1985) and for other loliginids by Haefner (1964). Indices prior
to treatments were observed to be heterogeneous between
lots. In order to overcome these constraints, differences be-
tween indices before and after each treatment were calculated

and covariance analysis (ANCOVA) was applied to the new
sets of variables using ML as covariate. Adjusted means and
95% confidence intervals were determined by the GT2
method, using the modification of Gabriel (Sokal and Rohlf,
1981). The overlap of confidence intervals between any pair
of treatments indicated whether or not they operate in
significantly different ways (Fig. 2).

Differences were found in FWI and EDI between the
groups placed directly into formalin vs. alcohol. Arm length
index, TLI and FLI of the lot fixed in ethanol differed from the
frozen and ethanol fixed lot. Fin length index, TLI and FWI
of the lot directly fixed in formalin differed from the previous-
ly frozen one. Only formalin fixation following freezing and
ethanol fixation following freezing did not show significant dif-
ferences in any of the calculated indices.

Except for tentacle and arm indices, most differences
between treatments were no more observed after long-term
preservation. No differences were detected between animals
preserved in formalin and in ethanol, as well as between those
previously frozen. However, material fixed and preserved
directly in ethanol was different from that preserved follow-
ing freezing in terms of ALI and TLI, and specimens fixed and
preserved directly in formalin differed from those previously
frozen in their indices of arm length, tentacle length and eye
diameter.

DISCUSSION

Many teuthologists have expressed concern about the
validity of comparing measurements and morphometric in-
dices of specimens of Loliginidae subjected to different fixa-
tion and preservation procedures (Haefner, 1964; LaRoe, 1967;
Cohen, 1976; Hixon et al., 1981). Haefner (1964) found arms
and tentacles to shrink more than 5% in Loligo pealei Lesueur,
1821 and Lolliguncula brevis (Blainville, 1823) preserved in 5%
formalin, and showed that growth of those species is allo-
metric, indicating that it is imprudent to compare indices from
groups having different sizes. LaRoe (1967) observed a con-
traction of 1.3% in ML of 15 specimens of Doryteuthis plei
(Blainville, 1823) fixed in 10% formalin and preserved in 70%
ethanol. Hixon et a/. (1981) point out an approximate 5%
shrinkage for Loligo pealei fixed in 10% formalin and later
transferred to 55% isopropanol.

As far as we know, only Cohen (1976) compared body
proportions in specimens submitted to different fixation pro-
cedures. She found differences in the adjusted means of
ANCOVA performed with ML as covariate for Loligo pealei

ANDRIGUETTO AND HAIMOVICI: PRESERVATION EFFECTS ON SQUID

MEASUREMENTS
ML
=
AL —
ee —
FL an
ED
I T I T
ML an
tl =
[— com: |
AL =a
| —— =]
FW Ll
FL ae
ED: =a
I T T
ao
ML |
| eet om |
aie |
| om om |
AL
co
FW
co
FL
=
ED
[ T T ]
ML o™ |
Tals ==
AL a=
FW |
FL ao 7
ED ==
i} T T
ML i
Te =
AL —
FW = |
FL =— |
ED = =
[ T T T T 1
07 08 09 10 1 1,2

formalin

ethanol

freezing

freezing - ethanol

iS
o
=
=
i)
'
aD
N
o
o
=

INDICES
—
ALI a Sas
FWI aa
FLI a
EDI ——
I T T T i}
Ua
| ee os |
ALI a=
FWI =
FLI an”
EDI —
T i T
TLI
co
ALI
oc
FWI
| oa
FLI
q
EDI
=
T T _ I l
TLI
ALI =a
FWI ==
FLI —_
EDI on 2
——
===
ar es | —\y eal ~ al
TE — |
ALI ] aa
Wi a |
FL a
EDI a
i T T t T 1
07 08 09 10 1 1,2

Fig. 1. Means and 95% confidence intervals of the quotients of measurements and indices by treatments. White bars indicate short-term changes;
black bars indicate long-term changes. A change is significant when a confidence interval does not include unity.

fixed in 10% formalin and preserved in 40% isopropyl alcohol,
when comparing lots of specimens fixed immediately after
capture with those previously refrigerated for 48 hours.

Our experiment included other treatments and periods
than those tested by Cohen. In addition, we compared
changes rather than absolute differences in measurements
and indices between lots. This enabled us to compare
heterogeneous lots.

Refrigeration and freezing are common methods for
stocking squids in commercial fishing, and are useful if fix-
atives, such as formalin are forbidden on board. The results
of Cohen (1976) and those presented here show that some
indices in previously refrigerated or frozen specimens are not
comparable with those of directly fixed ones, even if the same
chemicals were used. The same applies to preservation in
ethanol and formalin for several years, although to a lesser
degree.

Long-term preservation effects increased the mean
variance of indices and consequently the width of confidence
limits, making the discrimination of real differences from
preservation artifacts more difficult. Despite the small number
of indices included in our analysis, the results show that
numerical comparisons of populations or species of loliginids
based on body dimensions and proportions should consider
fixation and preservation induced artifacts, if lots were treated
in different ways. The measurement of just caught, fresh
specimens, or the comparison of lots fixed and preserved in
the same way for similar periods is advisable, unless effects
of specific treatments on indices are previously known.

ACKNOWLEDGMENTS

We wish to gratefully acknowledge J. R. Cure and H. Koehler
who assisted with computer calculations.

216

ARM LENGTH INDEX

ethanol

formalin

freezing

freezing —ethanol

freezing -formalin

FIN WIDTH INDEX

ethanol

formalin

treezing
freezing-ethanol

freezing -formalin

FIN LENGTH INDEX

ethanol
formalin
freezing
freezing-ethanol

freezing -formalin

EYE DIAMETER INDEX

ethanol

formalin

freezing
freezing-ethanol

treezing-formalin

TENTACLE LENGTH INDEX

ethanol
formalin
freezing
treezing-ethanol

freezing-formalin

AMER. MALAC. BULL. 6(2) (1988)

—,
ae
_—Sas en |
|
— |
a
———
[a aay
T a ] ] }
-0,06 -004 -0,02 0 002 004 006
<7
=e
[_ es Ee |
Se
SSS
ee
c——,
EE ee
a - <a"? ] i a : ] I ]
-0,06 -0,04 -0,02 0 0,02 004 0,06
— |
a
Ee ei
———
| a Se |
a ED
= i — OT =a) <= T ]
-0,06 -004 -002 O 0,02 0,04 0,06
[ee
| Seems eos |
en eS
o>
SS a
- al al eal) aa | T ] |
-0,06 -004 -0,02 oO 0,02 0.04 006
) A (Re |
=
[SD |
es a
——)
eee
————s
f T T T T I }
=05 -04 -03 =? -01 (0) O1 0,2 03

Fig. 2. Adjusted means and 95% confidence intervals of differences between morphometric indices before and after each treatment. White
bars indicate short-term differences; black barks indicate long-term differences. Treatments differ significantly when their confidence intervals

do not overlap.

LITERATURE CITED

Brakoniecki, T. F. 1984. A full description of Loligo sanpaulensis
Brakoniecki, 1984 and a redescription of Loligo gahi D’Orbigny,
1835, two species of squid (Cephalopoda, Myopsida) from the
Southwest Atlantic. Bulletin of Marine Science 34(3):435-448.
Castellanos, Z. J. A. 1967. Contribucion al estudio bioldgico de Loligo
brasiliensis Bl. Boletin del Instituto de Biologia Marina 14:5-35.
Cohen, A. C. 1976. The systematics and distribution of Loligo
(Cephalopoda, Myopsida) in the Western North Atlantic with
descriptions of two new species. Malacologia 15:299-367.
Figueiras, A. y O. E. Sicardi. 1980. Catalogo de los moluscos marinos
del Uruguay. Parte X: Revision actualizada de los moluscos
marinos del Uruguay con descripcion de las especies
agregadas. Seccion II: Gastropoda - Cephalopoda y

bibliografia consultada. Comunicaciones de la Sociedad
Malacologica del Uruguay V(38):179-277.

Haefner, P. A. 1964. Morphometry of the common Atlantic squid, Loligo
pealei, and the brief squid, Lolliguncula brevis, in Delaware Bay.
Chesapeake Science 5(3):138-144.

Haimovici, M. and J. M. Andriguetto Jr. 1986. Cephalopods in bot-
tom trawl fishing off south brazilian coast. Arquivos de Biologia
e Tecnologia do Instituto de Tecnologia do Parana 29(3):473-495.

Hixon, R. F., R. T. Hanlon and W. H. Hulet. 1981. Growth and max-
imal size of the long-finned squid Loligo pealei in the north-
western Gulf of Mexico. Journal of Shellfish Research
1(2):181-185.

Juanico, M. 1979. Contribuicao ao estudo da biologia dos
Cephalopoda Loliginidae do Atlantico Sul Ocidental, entre Rio

ANDRIGUETTO AND HAIMOVICI: PRESERVATION EFFECTS ON SQUID rad

de Janeiro e Mar del Plata. Doctoral Dissertation, Instituto
Oceanografico de Sao Paulo, Brazil. 102 pp.

Juanico, M. 1980. Developments in South American squid fisheries.
Marine Fisheries Review 42(7-8):10-14.

Kashiwada, J. and C. W. Recksiek. 1978. Possible morphological in-
dicators of population structure in the market squid, Loligo
opalescens. California Department of Fish and Game, Fish
Bulletin 169:99-111.

LaRoe, E. 1967. A contribution to the biology of the Loliginidae
(Cephalopoda: Myopsida) of the tropical western Atlantic.
Master’s Thesis, University of Miami, Florida. 220 pp.

Roper, C. F. E. and G. L. Voss. 1983. Guidelines for taxonomic descrip-
tions of cephalopod species. Memoirs of The National Museum
of Victoria 44:48-63.

Roper, C. F. E. and M. J. Sweeney. 1983. Techniques for fixation and

preservation of cephalopods. Memoirs of The National Museum
of Victoria 44:28-47.

Sokal, R. R. and F. J. Rohlf. 1981. Biometry, 2nd ed. W. H. Freeman,
New York. 859 pp.

Vigliano, P. H. 1985. Contribucion al conocimiento de la biologia de
Loligo brasiliensis Blainville, 1823 (Mollusca, Cephalopoda) en
aguas argentinas. Doctoral Dissertation, Facultad de Ciencias
Naturales y Museo de la Universidad Nacional de La Plata,
Argentina. 183 pp.

Voss, G. L. 1974. Loligo surinamensis, a new species of loliginid squid
(Cephalopoda, Myopsida) from northeastern South America.
Zoologische Mededelingen 48(6):43-53.

Voss, G. L. 1977. Present status and new trends in cephalopod
systematics. Symposium of the Zoological Society of London
38:49-60.

Date of manuscript acceptance: 15 September 1987.

INDEX TO THE AMERICAN MALACOLOGICAL BULLETIN: 1983 TO 1988

VOLUMES 1 THROUGH 6, SPECIAL EDITION
NUMBERS 1-3

CLEMENT L. COUNTS, III

Coastal Ecology Research Laboratory
University of Maryland Eastern Shore
Princess Anne, Maryland 21853, U.S.A.

With the appearance of Volume 6, No. 2, the American Malacological Bulletin completes its first six years of publication.
The Bulletin succeeded the Bulletin of the American Malacological Union in 1982 when it was recognized that an expanded
format was necessary to communicate the proceedings of the annual meetings of the American Malacological Union in a reviewed
format as well as to provide an outlet for malacological research not necessarily presented at A.M.U. meetings. Since the ap-
pearance of Volume 1 in 1983, the American Malacological Bulletin has published nine issues totaling 1,161 pages of primary
research articles (111 papers with a total of 1061 pages and 310 research abstracts with a total of 100 pages). Also, during
the first six years, the Bulletin has published three Special Editions comprising 47 research papers (428 pages) and 1 abstract
(1 page). Thus, the Bulletin has published 158 papers and 311 abstracts for a total of 1,590 pages during its first six years.

Because of this expanded publication format, it was felt that an index to the first six volumes and three special publica-
tions was necessary to increase their usefulness as a malacological research resource. Accordingly, the following index was
assembled to include authors, taxonomic groups, geographic localities, and various major subject headings. Each major category
is provided as a separate index.

DATES OF PUBLICATION AND KEY

The following is a compilation of the dates of publication of the first six volumes of the American Malacological Bulletin
and the first three Special Editions. The abbreviations for volume and issue numbers are in brackets following each date of
publication. These abbreviations are used throughout the index.

Volume 1: May 1983 [1]

Volume 2: February 1984 [2]

Volume 3, No. 1: December 1984 [3(1)]
Volume 3, No. 2: June 1985 [3(2)]
Volume 4, No. 1: February 1986 [4(1)]
Volume 4, No. 2: August 1986 [4(2)]
Volume 5, No. 1: January 1987 [5(1)]
Volume 5, No. 2: June 1987 [5(2)]
Volume 6, No. 1: January 1988 [6(1)]
Volume 6, No. 2: July 1988 [6(2)}
Special Edition No. 1: July 1985 [S1]
Special Edition No. 2: June 1986 [S2]
Special Edition No. 3: October 1986 [S3]

ACKNOWLEDGMENTS

| wish to thank Drs. Robert S. Prezant and Ronald B. Toll for their review and editing of this index. Thanks are also due Edward R.
Urban, Jr., who helped assemble the final draft of the manuscript.

219

220 AMER. MALAC. BULL. AUTHOR INDEX: 1983 - 1988

Abbe, George R.: S3:59-70

Adamkewicz, Laura: 1:107

Ahlstedt, S. A.: 1:43-50; 4(2):231

Al-Mousawi, Basima: 5(1):125-128

Albuquerque, B. L.: 2:97

Aldrich, Frederick A.: 2:51-56

Aldridge, David W.: 3(2):169-177

Amaratunga, Tissa: 1:90

Ambrose, R. F.: 2:90

Anderson, Roland: 4(2):241

Anderson, William D.: 3(1):102; 4(1):111

Andriguetto, Jr., José Milton: 6(2):213-217

Ashdown, M.: 1:103

Audesirk, Gerald: 2:78

Audesirk, T. E.: 2:78

Auffenberg, Garth: 3(1):98-99

Auffenberg, Kurt: 1:89; 3(1):98-99

Ayvazlan, Suzanne G.: 4(1):120

Babrakzai, Noorullah: 1:106; 2:97

Balboni-Tashiro, Jay Shiro: 4(1):118-119,
121-122; 4(2):236-237

Balch, N.: 4(1):55-60

Balch, Norval: 4(2):240-241

Banks, Glynn E.: S3:37-40

Bargar, Tom.: 3(1):83-84

Bates, John M.: 1-93

Beck, Malcolm L.: 1:97-98

Benamy, Elana: 3(1):92

Benjamin, Richard B.: 3(2)201-212

Bexerra, M. Z. B.: 1:67-70

Bieler, Rudiger: 4(1):108-109; 4(2):236

Blum, Bernard J.: 3(1):92

Bogan, Arthur E.: 1:93-94, 98; 3(1):1-10,
105-106; 4(1):25-37; 6(1):19-37

Boletzky, Sigurd V.: 4(2):217-227

Boss, Kenneth J.: 4(2):236

Boucher-Rodoni, R.: 4(2):240

Bouchet, Philippe: 4(1):49-54

Bowman, Charles F.: S2:95-98

Bowser, Amy: 4(1):121-122

Bradford, Lea A.: 2:93

Bronmark, Christer: 5(1):73-84

Brown, Kenneth M.: 3(2):143-150;
5(1)73-84; 6(1):9-17

Bublitz, C. G.: 2:89

Buchanan, Alan C.: 2:85; 4(1):119

Buckley, Daniel E.: 3(2):268

Buckley, George D.: 2:96

Bullock, Robert C.: 4(1)114-115

Burch, Beatrice L.: 2:83

Burch, J. B.: 2:88-89

Burch, Thomas A.: 2:83

Burky, Albert J.: 3(1):94; 3(2):135-142,
201-212

Buroker, Norman E.: 1:108

Buttner, Joseph K.: S2:211-218

Cain, Arthur J.: 1:105-106; 2:75-76, 82

Cairns, John, Jr.: 4(1):116; S2:69-81

Call, Samuel M.: 1:31-34

Calvo, lara S.: 3(1):101-102

Camburn, Keith E.: 3(1):47-53

Cameron, Robert: 1:103

AUTHOR INDEX

Campbell, John H.: 4(2):242

Campbell, Lyle D.: 3(1):96; 4(1):39-42

Campbell, Sarahlu C.: 4(1):39-42

Carriker, Melbourne R.: 1:35-42, 102;
2:75-76; 4(1):119; S3:41-49, 71-74

Carter, M. A.: 1:103

Carter, W. R., Ill: S3:5-10

Chabot, Jennifer: 4(2):236-237

Chalermwat, Kashane: 2:87; 3(1):101;
4(1):115-116

Chamberlain, J. A., Jr: 4(2):239-240

Chambers, Steven M.: 1:109

Chen, Deli: 2:88

Chen, Pulin: 2:88

Cherry, Donald S.: 4(1):116; S2:69-81

Chrisman, C. Larry: 1:106-107

Christensen, Carl C.: 1:97; 2:98-99

Cicerello, Ronald R.: 3(1):47-53

Clark, Kerry B.: 5(2):259-280

Clarke, Arthur H.: 1:27-30; 3(1):104-105

Cloney, Richard A.: 2:91

Coan, Eugene: 1:89; 2:83; 3(1):103

Coelho, M. L.: 4(2):239

Cohen, George: 4(1):121-122

Cookson, Ellen: 3(1):89-90

Coney, C. Clifford: 1:94-95, 95

Conover, Denis G.: 3(2):201-212

Cooper, Kay M.: 2:91, 93-94

Cordoba, Eileen: 4(2):231-232

Counts, Clement L., Ill: 1:100; 4(1):81-88;
4(2):230; S2:7-39

Covich, Alan P.: 5(1):73-84

Cowie, Robert H.: 1:104

Cox, Carollyn: 5(1):49-64

Craveiro, A. A.: 1:67-70

Crawford, Maurice K.: 4(1):120-121

Creitz, Michael R.: 1:95

Croz, L. D.: 4(1):119

Culter, James K.: 4(1):107

Cummins, H.: 1:89

Daiber, Franklin C.: S1:iii

D’Asaro, Charles N.: 4(2):185-199

Davis, George M.: 1:109-110; 2:75-76,
88; 3(1):96

DeFreese, Duane: 5(2):259-280

Deisler, Jane E.: 2:98; 3(1):103

Deitz, Thomas H.: 3(2):233-242

DeLancey, L. B.: 4(2):240

Dennis, Sally: 1:93

Denny, M. W.: 4(2):242-243

Dermot, M. Edwin: 2:90

DeRusha, Randal H.: 2:92-93

Dexter, Ralph W.: 4(1):112-113

Deyrup-Olsen, |.: 2:91

Diaz-Tous, |. A.: S2:83-88

Dillon, Patrick: 1:103

Dillon, Robert T.: 1:105; 3(1):99-100;
5(1):101-104

DiNuzzo, Anthony R.: 2:93-94

Doherty, F. G.: 4(1):116

Dougherty, B. J.: 3(1):99

Downing, Gary G.: $2:185

Draper, Bertram C.: 4(2):232-233

Dunhardt, Patricia A.: S2:69-81

Dussart, G. B. J.: 5(1):65-72

Earhart, H. Glenn: S3:11-16

Ebert, Danny: 4(1):21-23

Edmunds, Malcolm: 5(2):185-196

Eernisse, Douglas J.: 4(2):243

Eisensamer, Brigitte: 6(1):131-139

Eldridge, Peter J.: 2:96-97; 4(2):149-155

Emberton, Kenneth C.: 1:98; 2:97-98

Emerson, William K.: 1:75-76

Esposito, Mark A.: 3(2):179-186

Estes, James A.: 2:80

Etter, Ron J.: 4(1):110

Eversole, Arnold G.: 2:96-97; 3(1):102;
4(1):111; 4(2):149-155

Ewart, John W.: 4(1):119

Eyster, Linda S.: 4(2):205-216

Fairbanks, H. L.: 4(2):238

Fairbanks, H. Lee: 1:21-26

Fallo, Glen J.: 3(1):47-53

Farache, Vivianne: 1:92

Fields, Patrick F.: 2:85

Fischer, Franz Peter: 6(1):131-139, 153-159

Flessa, Karl W.: 2:79-80

Foe, Christopher: S2:133-142, 143-150

Folse, Dean S.: 2:93-94

Foltz, David W.: 1:109-110

Forsythe, John W.: 2:92, 92-93, 93-94

Foy, E. A.: 4(1):55-60

Fraley, N.: 4(2):241

Franklin, Dee A.: 1:106-107

Freitag, Thomas M.: 3(1):105

Fritz, Lowell W.: 3(1):100-101

Fukuyama, Alan: 1:91-92

Fukuyama, Alan K.: 2:94

Fuller, S. Cynthia: 4(2):233-234

Galloway, Marvin L.: 4(1):61-79, 116;
$2:193-201

Garton, David W.: 2:63-73

Gee, Penelope A.: 2:94

Gilly, W. F.: 4(2):241

Goldman, Michael A.: 1:106-107

Gomez, J. A.: 4(1):119

Goodfriend, Glenn A.: 1:99-100

Gordon, Mark E.: 1:97; 3(1):100;
4(1):115, 116

Gosline, J. M.: 2:90

Gosliner, Terrence M.: 2:95-96;
5(2):243-258

Gosling, Elizabeth: 1:108

Green, R. H.: 1:90, 108-109

Greenwood, Jeremy J. D.: 1:103

Grimes, Lawrence W.: 2:96-97;
4(2):149-155

Gustafson, Richard: 2:94

Haas, Dieter: 5(1):85-90

Hadfield, Michael G.: 5(2):197-214

Haimovici, Manuel: 6(2):213-217

Hall, James J.: 1:96

Han, Jonathan Kyung Ho: 4(1):118-119

Hanley, Robert W.: 1:94; 2:87-88

AMER. MALAC. BULL. AUTHOR INDEX: 1983 - 1988

Hanlon, Roger T.: 2:91, 92, 92-93, 93, 93-94

Harasewych, M. G.: 3(1):11-26; 4(2):233

Hargreave, David: 4(1):108

Harris, Larry G.: 5(2):287-292

Harry, Harold W.: 1:90; 4(2):157-162

Hartfield, Paul: 4(1):21-23

Harvell, C. Drew: 2:83-84

Haven, Dexter S.: S3:17-23

Havenhand, Jonathan D.: 4(1):103-104

Havlik, Marian E.: 1:51-60; 3(1):106-107;
4(2):230, 230-231

Hay, William: 1:99

Hayes, D. R.: 4(1):110-111

Hayes, P. F.: S2:41-45, 47-52

Heard, W. H.: 4(1):101

Hedgecock, Dennis: 1:108

Heller, Joseph: 1:104

Helm, P. L.: 4(1):55-60

Henager, C. H.: S2:47-52

Hendrickson, Lisa C.: 4(1):110

Hendrix, Serman S.: 4(1):119

Hershler, R.: 4(2):243

Hickman, Carole S.: 3(1):95; 4(1):114;
4(2):242

Hicks, B.: 1:90

Hill, David M.: 5(2):153-157

Hillman, Robert E.: $1:101-109

Hixon, Raymond F.: 2:93

Hoagland, K. Elaine: 1:110; 2:88;
3(1):33-40, 85-88; 4(1):88-99;
4(2):173-183; S2:203-209

Hochberg, F. G.: 2:98

Hoeh, Walter R.: 3(1):92-93; 4(2):231-232

Hoffman, J. E.: 4(1):113-114

Hoggarth, Michael A.: 2:82, 86;
4(1):117-118

Hoke, Ellet: 1:71-74

Holland-Bartels, L. E.: 6(1):39-43

Holopainen, Ismo J.: 5(1):21-30, 41-48

Horn, Karen J.: 1:61-68; 2:86

Hornbach, Daniel J.: 3(2):187-200;
5(1):49-64

Horrigan, F.: 4(2):241

Houbrick, Richard S.: 2:1-20; 3(1):96;
4(1):109; 4(2):235

Houck, B. A.: 2:90-91

Hunter, Margaret A.: 6(1):1-8

Hurley, Geoffrey V.: 4(2):240-241

Imlay, Marc J.: 1:97; 3(1):107

Isom, Billy G.: 1:93; S2:1-5, 95-98

Jablonski, David: 2:79-80; 4(1):49-54

James, Frances C.: 1:95

James, Matthew J.: 2:80-81, 85; 3(1):98

Jefferts, K.: 4(2):241

Jenkinson, John J.: 2:86-87

Johnson, Joseph T.: S2:95-98

Johnson, K. |.: S2:47-52

Jokinen, Eileen: 3(1):99; 5(1):9-19

Jonas, M.: 4(2):232

Kaas, Piet: 6(1):115-130

Kabat, Alan R.: 2:94

Kat, P. W.: 4(1):107

Kelly, Michael T.: 2:93-94

Kempf, S. C.: 4(2):235

Kennedy, George L.: 4(2):238

Kennedy, V. S.: 4(1):101

Kennedy, Victor S.: $3:25-29

Kent, Brett, W.: 2:79

King, Christina A.: 4(1):81-88

Kitchel, Helen E.: 3(1):104

Klippel, Walter E.: 3(1):41-44

Klosiewski, Steven P.: 5(1):73-84

Knight, Allen: S2:133-142, 143-150

Kohn, Alan J.: 2:81; 3(1):95; 4(2):236

Kool, Silvard P.: 1:94-95; 4(1):110, 4(2):233

Kotrla, M. Bowie: 1:95; 3(1):99; 4(1):117;
4(2):231

Kraemer, Louise Russert: 1:13-20, 83-88;
2:87; 4(1):61-79, 116; S2:187-191,
193-201

Kraemer, Robert: S2:193-201

Krejci, Mark J.: 2:93

Kubodera, Tsunemi: 2:89-90

Kuo, Yuanhua: 2:88; 3(1):96

Lacey, Will H.: 4(1):111

Lane, Roger L.: 3(1):27-32

Lang, M. A.: 4(2):241-242

Langdon, Christopher J.: 4(1):81-88

LaRochelle, Peter B.: 1:99

Lasalle, Mark W.: S3:31-36, 71-74

Lauritsen, Diane D.: 3(1):101; S2:219-222

Leathers, Bonnie K.: 5(1):73-84

Lechleitner, Richard A.: S2:69-81

Leise, Esther M.: 6(1):141-151

Lera, Monica: 1:92

Lietzow, Jeffrey S.: S2:185

Lillico, Stuart: 2:81

Lindberg, David R.: 2:80, 95; 4(1):115;
4(2):244

Linden, Lawrence H.: S2:53-58

Little, Colin: 3(2):223-231

Lloyd, Philip: 2:78

Lodge, David M.: 5(1):73-84

Loomis, S. H.: 4(1):110-111

Lopez, Glenn R.: 5(1):21-30

Lord, Acha: 4(2):201-203

Loverde, Philip T.: 1:106-107

Lu, C. C.: 4(1):101

Lunz, John D.: S3:31-36

Lutz, Richard A.: 1:101; 3(1):100-101;
4(1):49-54; S1:59-78

Lyons, William G.: 1:91; 3(1):97-98;
6(1):79-114

Machado, M. I. L.: 1:67-70

Mackie, Gerald, L.: 4(1):116; 5(1):31-39;
$2:223-229

MacPhee, D. David: S2:59-61

Maddox, Nora V.: 4(1):107

Mahieu, Genoveva C. de: 1:92

Malek, E. A.: 1:67-70

Mangold, K.: 4(2):240

Mann, Roger: S3:51-57, 71-74

Marcus, Eveline Du Bois-Reymond:
5(2):183-184

Marking, Leif L.: 3(1):106-107

Martin, A. W.: 2:91

221

Mattice, J. S.: S2:167-178

Mazurkiewicz, Michael: 4(1):101-102

McCuaig, J.: 1:90

McKee, Susan J.: 4(2):237

McLean, James H.: 2:21-34; 3(1):104;
4(1):109

McLeod, Michael J.: 1:96; S2:125-132

McMahon, Robert F.: 3(2):135-142,
243-265, 267-269; 5(1):105-124;
$2:99-111, 151-166, 231-239

McNair, E. C., Jr.: S3:37-40

Meier-Brook, Claus: 5(1):85-90

Merrill, Arthur S.: 4(2):236

Messenger, J. B.: 2:92

Metcalf, Art L.: 2:86

Mikkelsen, Paula M.: 1:91; 3(1):93;
4(2):233

Mikkelsen, Paul S.: 1:91, 100; 3(1):93,
93-94; 4(2):233

Millen, Sandra V.: 2:95

Miller, Andrew C.: 5(2):177-179; 6(1):49-54

Miller, Stephen E.: 5(2):197-214

Miller, Walter B.: 1:21-26, 106; 2:97, 98

Miles, Charles D.: 1:97-98

Miltz, Christina: 6(1):131-139

Mitchell, L. G.: 6(1):39-43

Moore, Donald R.: 1:89; 3(1):103-104

Moore, Richard H.: 1:94-95, 95

Morton, Brian: 4(2):233; 5(1):91-99;
5(2):159-164; S2:113-124

Morris, Claude C.: 2:51-56

Morse, M. Patricia: 2:95; 5(2):281-286

Moyer, Steven N.: 3(1):106; 6(2):179-188

Muldoon, Kathryn A.: 3(1):93

Muldoon-McLaughlin, Kathryn: 1:100

Mulvey, Margaret: 1:107

Murray, Harold D.: 1:95-96

Murray, James: 1:103-104, 104

Mussalli, Yusuf G.: S2:83-88

Nash, Kelly L. (Clayton): $2:185

Neck, Raymond W.: 1:99; 2:86;
S2:179-184

Neitzel, D. A.: S2:41-45

Neves, Richard J.: 5(1):1-7; 6(2):179-188

Newball, Sara: 1:35-42

Nishiyama, T.: 2:89

Nutall, T. R.: 4(2):232

Nybakken, James: 1:91-92

O’Dor, R. K.: 3(1):107; 4(1):55-60

Oliveira, G. P: 2:97

Page, T. L.: S2:41-45, 47-52

Palmer, A. Richard: 1:105

Parkin, David T.: 1:103

Parmalee, Paul W.: 3(1):41-44; 4(1):25-37;
6(2):165-178

Parsons, A. Michelle: 2:93

Paulay, Gustav: 2:83

Payne, Barry S.: 5(2):177-179; 6(1):49-54

Pearce, Timothy A.: 3(1):98; 4(2):237

Pearcy, W. G.: 4(2):241

Pechenik, Jan. A.: 4(2):165-172; S1:85-91

Penchaszadeh, Pablo E.: 1:92

Perron, Frank E.: 4(2):229

222 AMER. MALAC. BULL. AUTHOR INDEX: 1983 - 1988

Peters, Gregory T.: S2:69-81

Petit, Richard E.: 1:79-80; 2:57-61

Petuch, Edward J.: 2:79

Poizat, Claude: 5(2):303-306

Porter, Hugh J.: 1:61-68; 3(1):100;
4(1):107-108

Potter, Jeanne Miles: S2:53-58

Powell, E. N.: 1:89

Prezant, Robert S.: 1:101-102, 102;
2:41-50, 87; 3(1):104; 4(1):116-117;
4(2):235; 5(2):173-176; S1:35-50;
$3:1-4

Pritchard, Donald W.: S3:71-74

Purser, G. John: 5(1):125-128

Quinn, James F., Jr: 1:92; 2:84; 3(1):97-98

Raeihle, Dorothy: 1:75-77

Rajasekaran, S.: 4(1):114; 4(2):237

Reed-Miller, Charlene: 1:102

Reeder, Richard L.: 1:96-97, 98; 2:98;
4(2):237

Reid, David G.: 4(1):112

Reid, R. G. B.: 2:83

Richards, Charles S.: 1:106

Richardson, Terry D.: 6(1):9-17

Rios, Eliezer de Carvalho: 1:92; 2:97;
3(1):101-102; 4(2):233

Rittschof, Dan: S1:111-116

Rivest, Brian R.: 4(2):229

Robertson, Robert: 1:1-12; 4(1):113;
$1:1-22

Robinson, Kenneth: 1:31-34

Rodgers, Elizabeth B.: S2:95-98

Rogers, Steffen H.: 1:96-97, 98;
3(1):89-90

Rogge, Thomas N.: 4(1):111; 4(2):234

Roller, Richard A.: 2:63-73; 6(2):189-197

Rollins, Harold B.: 3(1):96-97

Rollinson, D.: 1:107

Roper, Clyde F. E.: 2:89; 3(1):55-61,
63-82; 4(1):101; S1:93-100

Ropes, John W.: 4(1):120-121

Rosenberg, Gary: 2:84

Rosenfield, David S.: 5(2):153-157

Rosewater, Joseph: 1:90-91; 2:35-40;
3(1):107

Roth, Barry: 1:98; 2:98; 3(1):1-10, 102-103

Rouquayrol, M. Z.: 1:67-70

Roy, Rob L.: S2:69-81

Russell-Hunter, W. D.: 3(2):213-221,
269-272; 6(1):69-78

Sacchi, Cesare F.: 1:107-108

Sanchez, Modesto, Jr.: 5(2):153-157

Saul, L. R.: 4(2):236

Scheltema, Amelie H.: 3(1):97; 6(1):57-68

Schick, Daniel F.: 6(1):1-8

Schmidt, John E.: 4(1):117

Scott, Paul H.: 2:96; 4(2):234

Scott-Wasilk, Jennifer: S2:185

Seeley, Robin Hadlock: 1:92; 4(1):108

Selander, Robert K.: 1:110

Shasky, Donald R.: 2:84

Shimek, Ronald: 2:82, 91-92, 94-95

Shumway, Sandra E.: 6(1):1-8

Sickel, James B.: S2:83-88, 89-94

Sigel, Liz: 4(1):121-122

Sigurdsson, John Baldur: 4(1):101-103

Simmons, M. A.: S2:41-45

Sinclair, Ralph M.: 1:93

Singer, Ingrid: 6(1):131-139

Singh, S. M.: 1:90, 108-109

Sirois, Andre: 4(2):240-241

Smith, Douglas G.: 4(1):13-19

Smith, Judith Terry: 2:84-85; 4(1):1-12;
4(2):238-239

Smithson, James A.: S2:63-67

Snyder, S.: 4(2):241

Solem, Alan: 1:98-99; 2:97

Soliman, Gamil N.: 4(1):103, 109-110

Sorenson, Fred: 2:80

Sridharan, T.: 4(1):114

Sriramulu, Vijayam: 4(1):114; 4(2):237

Staff, G.: 1:89

Stansbery, David H.: 1:93; 2:86

Stanton, R. J., Jr: 1:89

Starnes, Lynn B.: 1:93-94; 3(1):105-106;
6(1):19-37

Starr, R. M.: 4(2):239

Stein, Roy A.: 5(1):73-84

Stickle, William B.: 2:63-73; 6(2):189-197

Strayer, D.: 4(1):119-120

Streit, Bruno: 3(2):151-168

Summers, William C.: 2:90

Swann, Charles P.: 1:102

Swanson, Charles: S2:193-201

Sweeney, Michael J.: 2:89; 3(1):63-82;
4(1):101

Tan Tiu, Antonieto: 3(1):103; 4(1):112,
116-117; 4(2):234; 5(2):173-176;
6(2):199-206

Tashiro, Jay Shiro: 3(2):179-186

Taub, Stephan R.: 1:107

Taylor, Ralph W.: 2:85-86

Theler, James L.: 5(2):165-171

Thiriot-Quievreux, Catherine: 1:105-106

Thorsson, Wesley: 2:81

Tissot, B. N.: 4(2):234-235

Todd, C. D.: 4(2):235

Todd, Christopher D.: 4(1):103;
5(2):293-301

Toll, Ronald B.: 2:89; 6(2):207-211

Topping, Jane M.: 2:82

Torelli, Alberto A.: 1:92-93

Trdan, Richard J.: 3(1):92-93;
4(2):231-232

Tremblay, M. J. 4(1):104

Tripp, Marenes R.: S$1:79-83

Turk, Philip E.: 2:93

Turner, Ruth D.: 3(1):95-96; 4(1):49-54;
$1:23-24

Van Belle, Richard A.: 6(1):115-130

Van Der Schalie, Henry: 1:93

Van Heukelem, W.: 4(1):101

Vecchione, Michael: 1:90; 4(1):45-48, 101

Vermeij, Geerat J.: 2:79

Villalaz, Janzel A.: 4(1):119

Villoch, Margarita R.: 2:93

Virnstein, Robert W.: 3(1):93-94

Voight, Janet R.: 6(1):45-48

Voltzow, Janice: 4(1):110; 4(2):243

Vrijenhoek, Robert C.: 1:107

Walborn, Patricia: 4(1):121-122

Wall, J. R.: 1:107

Waller, D. L.: 6(1):39-43

Waller, Thomas R.: 1:101; 4(1):111-112

Walsh, Lyle: 1:102

Ward, J. E.: 4(1):122

Ward, J. Evan: 3(1):97

Ward, Peter: 2:79, 90, 91

Waren, Anders: 2:83; 4(1):49-54

Warheit, Kenneth I.: 2:80

Warren, Melvin L., Jr.: 3(1):47-53

Way, Carl M.: 3(1):100

Webber, D. M.: 3(1):107

Wells, Fred E.: 3(1):97

Whitaker, J. D.: 4(2):240

Whitcomb, James P.: S3:17-23

White, Patricia A.: 3(1):94

Whitehead, Bruce E.: 5(1):105-124

Widlak, James C.: 3(1):106; 5(1):1-7

Wieland, Steven J.: 2:78

Wilbur, Karl M.: S1:51-58

Willan, R. C.: 5(2):215-241

Williams, Carol J.: 3(2):267-269; S2:99-111,
151-166, 231-239

Williams, James D.: 3(1)105-106

Winter, Gabriele: 5(1):85-90

Wolfe, Douglas A.: 3(1):94

Wright, C. A.: 1:107

Wright, L. L.: S2:167-178

Wu, Shi-Kuei: 1:96

Yang, Hongmu: 2:88

Yang, Won Tack: 2:93

Young, Mark: 5(1):125-128

Zeller, Traudel: 5(1):85-90

Zouros, E.: 1:109

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 223

Abra alba (Wood, 1802): 5(1):21-30 (passim)
Abralia astrolineata Berry, 1914: 3(1):63-82
Abralia astrostica Berry, 1904: 3(1):63-82
Abralia trigonura Berry, 1913: 3(1):63-82
Abraliopsis scintillans Berry, 1911: 3(1):63-82
Acado Commercon, 1792: 5(2):215-241
Acanthina tyrianthina Berry, 1957: 3(1):63-82
Acanthochites hemphilli (Pilsbry, 1893): 1:91
Acanthochites pygmaeus (Pilsbry, 1893): 1:91
Acanthochites rhodeus (Pilsbry, 1893): 1:91
Acanthochites rhodeus Pilsbry, 1893:
6(1):79-114
Acanthochites spiculosus Dall, 1889:
6(1):79-114
Acanthochites (Cryptoconchus) floridanus
(Dall, 1889): 6(1):79-114
Acanthochiton astriger (Reeve, 1847):

6(1):79-114

Acanthochiton pygmaeus (Pilsbry, 1893):
6(1):79-114

Acanthochiton spiculosus Dall, 1889:
6(1):79-114

Acanthochitona Gray, 1821: 6(1):115-130

Acanthochitona andersoni Watters, 1981:
1:91; 6(1):79-114

Acanthochitona ashbyi Leloup, 1937:
6(1):115-130

Acanthochitona astrigera (Reeve, 1847):
1:91; 6(1):79-114

Acanthochitona balesae Abbott, 1954:
1:91; 6(1):79-114

Acanthochitona bonairensis Kaas, 1972:
1:91; 6(1):79-114

Acanthochitona brunoi Righi, 1971:
6(1):79-114

Acanthochitona ciroi Righi, 1971:
6(1):79-114

Acanthochitona communis Risso, 1826:
1:91; 6(1):79-114

Acanthochitona crinita (Pennant): 6(1):69-78

Acanthochitona elongata Kaas, 1972:
6(1):79-114

Acanthochitona fascicularis (Linné, 1767):
6(1):79-114, 131-139, 141-151, 153-159

Acanthochitona ferreirai Lyons, 1988, sp.
nov.: 6(1):85-86

Acanthochitona hemphilli (Pilsbry, 1893):
1:91; 6(1):79-114

Acanthochitona hirudiniformis (Sowerby,
1832): 1:91; 6(1):79-114

Acanthochitona interfissa Kaas, 1972:
1:91; 6(1):79-114

Acanthochitona limbata Kaas, 1986:
6(1):115-130

Acanthochitona lineata Lyons, 1988, sp.
nov.; 6(1):90-92

Acanthochitona mahensis Winckworth,
1927: 6(1):115-130

Acanthochitona minuta (Leloup, 1980):
6(1):79-114

TAXONOMIC INDEX

Acanthochitona pygmaea (Pilsbry, 1893):
1:91; 6(1):79-114

Acanthochitona rhodea (Pilsbry, 1893):
1:91; 6(1):79-114

Acanthochitona roseojugum Lyons, 1988,
sp. nov.: 6(1):98-100

Acanthochitona saundersi: 6(1):69-78

Acanthochitona spiculosa (Reeve, 1847):
1:91; 6(1):79-114

Acanthochitona tabogensis Smith, 1961:
6(1):79-114

Acanthochitona terezae Guerra Junior,
1983: 6(1):79-114

Acanthochitona venezuelana Lyons, 1988,
sp. nov.: 6(1):96-98

Acanthochitona viridis (Pease, 1872):
6(1):79-114

Acanthochitona woodwardi Kaas and Van
Belle, 1988, sp. nov.: 6(1):126-127

Acanthochitona worsfoldi Lyons, 1988, sp.
nov.: 6(1):92-94

Acanthochitona zebra Lyons, 1988, sp.
nov.: 6(1):105-107

Acanthochitona (Notoplax) hemphilli
(Pilsbry, 1893): 6(1):79-114

Acanthochitones spiculosus (Reeve,
1847): 6(1):79-114

Acanthochitones spiculosus astriger
(Reeve, 1847): 6(1):79-114

Acanthochitonidae Pilsbry, 1893:
6(1):79-114; 6(1):115-130

Acanthochitonina Bergenhayn, 1930:
6(1):115-130

Acanthochitoninae Ashby, 1925:
6(1):115-130

Acanthodoris Gray, 1850: 5(2):243-258

Acanthodoris brunnea MacFarland, 1905:
5(2):197-214

Acanthodoris hudsoni MacFarland, 1905:
5(2):197-214

Acanthodoris nanaimoensis O'Donoghue,
1921: 5(2):197-214

Acanthodoris pilosa (Muller, 1776):
5(2):197-214

Acanthopleura Guilding, 1829: 4(1):114-115;
6(1):115-130

Acanthopleura brevispinosa (Sowerby,
1840): 6(1):115-130

Acanthopleura gemmata (Blainville):
6(1):115-130

Acanthopleura granulata (Gmelin, 1791):
4(1):114-115; 6(1):79-114; S1:1-22

Acanthopleura haddoni Winckworth, 1927:
6(1):115-130

Acanthopleura spiniger: 6(1):115-130

Acanthopleura vailantii Rochebrune, 1882:
6(1):115-130

Acanthopleurinae Dall, 1889: 6(1):115-130

Acanthophora spicifera (Vahl) Bogesen:
5(2):259-280 (passim)

Acanthotrophon sentus Berry, 1969:
3(1):63-82

Acantopleura (sic) vaillantii Rochebrune,
1882: 6(1):115-130

Achatina fulica Bowditch: 2:98-99; 6(1):16

Achatinellidae: 4(1):112-113

Aciculidae: 3(2):223-231

Aclididae: $1:1-22

Aclis Lovén, 1846: $1:1-22

Acmaea acutapex Berry, 1960: 3(1):63-82

Acmaea concreta Berry, 1963: 3(1):63-82

Acmaea gabatella Berry, 1960: 3(1):63-82

Acmaea goodmani Berry, 1960: 3(1):63-82

Acmaea lepisma Berry, 1940: 3(1):63-82

Acmaea stanfordiana Berry, 1957:
3(1):63-82

Acmaea scabra Gould, 1846: S1:35-50

Acmaea testudinalis (Muller, 1776):
6(1):69-78

Acmaeidae Carpenter, 1857: 2:95; 4(1):115

Acochlidiacea Kuthe, 1935: 2:95;
5(2):281-286; S1:1-22

Acropora palmata (Lamarck, 1816): 1:1-12

Acroteuthis Berry, 1913: 3(1):63-82

Acruroteuthis Berry, 1920: 3(1):63-82

Actaeon (Microglyphia) schencki Berry,
1957: 3(1):63-82

Acteocina Gray, 1847: 4(1):39-42

Acteocina sp.: 3(1):93, 98; 4(2):233; $1:1-22

Acteocina canaliculata (Say, 1822):
4(1):39-42; 5(2):197-214

Acteocina candei (Orbigny, 1842): 1:91;
3(1):93, 98

Acteocina lepta Woodring, 1928: 3(1):93, 98

Acteocina smithi (Bartsch, 1915):
5(2):243-258

Acteocinidae Pilsbry, 1921: 4(2):233;
$1:1-22

Acteon Montfort, 1810: 5(2):185-196; S1:1-22

Acteon flammeus (Gmelin, 1791):
5(2):243-258

Acteon fortis Thiele, 1925: 5(2):243-258

Acteon tornatilis (Linné, 1758): 5(2):185-196

Acteon wetherilli Lea, 1833: 4(1):39-42

Acteonia cocksi Alder and Hancock:
4(2):205-216 (passim); 5(2):197-214

Acteonidae Orbigny, 1842: 5(2):243-258

Actinia equina Linné, 1758: 5(2):185-196

Actinonaias carinata (Barnes, 1823): 1:29,
43-50; 3(1):105; 6(1):19-37

Actinonaias carinata gibba (Simpson,
1900): 6(1):19-37

Actinonaias ellipsiformis (Conrad, 1836):
3(1):93

Actinonaias ligamentina (Lamarck, 1819):
3(1):41-45; 4(1):25-37; 6(1):19-37;
6(2):165-178

Actinonaias ligamentina carinata (Barnes,
1823): 1:31-34, 51-60; 2:85-86;
5(2):165-171

224 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Actinonaias pectorosa (Lea, 1827):
1:43-50; 3(1):104

Actinonaias pectorosa (Conrad, 1834):
6(1):19-37

Aculifera: 6(1):57-68

Adalaria Bergh, 1879: 5(2):197-214, 293-301

Adalaria lovéni (Adler and Hancock,
1862): 2:95

Adalaria pacifica Bergh, 1880: 2:95

Adalaria proxima (Adler and Hancock,
1854): 2:95; 4(1):103-104; 4(2)235;
5(2):197-214, 293-301; 6(1):17

Adamete viridula (Fabricius, 1780):
2:57-61

Adelomelon brasiliana (Lamarck, 1811):
4(2):165-172

Adenopod: 6(1):57-68

Adipicola Dautzenberg, 1927: S1:23-34

Admetula Cossmann, 1889: 2:57-61

Admetula evulsa (Solander, 1766): 2:57-61

Adontorhina Berry, 1947: 2:96; 3(1):63-82

Adontorhina cyclia Berry, 1947: 2:96;
3(1):63-82

Adula falcata (Gould, 1851): 5(2):159-164
(passim)

Aegries Loven, 1844: 5(2):243-258

Aegires albopunctatus MacFarland, 1905:
5(2):197-214

Aegires punctilucens (Orbigny, 1837):
5(2):197-214

Aegires sublaevis Odhner, 1932:
5(2):185-196, 197-214

Aeolidacea Orbigny, 1837: 4(2):205-216
(passim); 5(2):215-241

Aeolidia papillosa (Linné, 1761):
4(2):205-216; 5(2):185-196, 293-301;
6(1):57-68

Aeolidiella alba Risbec, 1928: 5(2):243-258

Aeolidiella alderi (Cocks, 1852):
5(2):303-306

Aeolidiella glauca (Alder and Hancock,
1845): 5(2):185-196

Aeolidiella indica Bergh, 1888: 2:95-96;
5(2):243-258

Aeolidiella sanguinea (Norman, 1877):
5(2):185-196, 303-306

Aeolidiidae Orbigny, 1837: 5(2):243-258

Aeolidiopsis Pruvot-Fol, 1956: 5(2):185-196

Aequipectin circularis (Sowerby, 1835):
4(1):119

Aequipecten (Leptopecten) camarella
Berry, 1968: 3(1):63-82

Aeromonas caviae: 2:82

Aforia circinata (Dall, 1873): 2:82

Agaronia murrha Berry, 1953: 3(1):63-82

Aglaja Renier, 1804: S1:1-22

Aglaja ocelligera (Bergh, 1894):
5(2):197-214

Aglajidae: 4(2):233; 5(2):185-196, 243-258;
$1:1-22

Akera Muller, 1776: S1:1-22

Akera soluta (Gmelin, 1791): 5(2):243-258

Akeridae Pilsbry, 1893: 5(2):243-258; $1:1-22

Alaba H. and A. Adams, 1853: 4(2):235

Alasmidonta Say, 1818: 6(2):165-178

Alasmidonta atropurpura (Rafinesque,
1831): 6(1):19-37

Alasmidonta calceolus (Lea, 1830): 1:43-50

Alasmidonta marginata Say, 1819: 1:43-50,
51-60; 3(1):104, 105; 4(1):117-118;
5(2):165-171; 6(1):19-37; 6(2):165-178

Alasmidonta minor (Lea, 1845): 1:43-50;
3(1):104; 6(1):19-37

Alasmidonta raveneliana (Lea, 1834):
6(1):19-37

Alasmidonta viridis Rafinesque, 1831:
1:29; 3(1):105; 4(1):117-118; 5(1):1-7;
5(2):165-171; 6(1):19-37; 6(2):165-178

Alasmidonta (Pressodonta) minor Lea,
1845: 6(2):165-178

Alba goniochila: 4(2):235

Alcyonium digitatum (Linne, 1758):
5(2):197-214

Alderia modesta (Loven, 1844):
5(2):197-214

Aldisa Bergh, 1878: 5(2):185-196

Aldisa banyulensis Pruvot-Fol, 1951:
5(2):185-196

Aldisa benguela ‘Gosliner’ Millen and
Gosliner, 1985: 5(2):243-258

Aldisa binotata Pruvot-Fol, 1953:
5(2):197-214

Aldisa cooperi Robilliard and Baba:
5(2):197-214

Aldisa pikokai Bertsch and Johnson:
5(2):197-214

Aldisa sanguinea (Cooper, 1862):
5(2):197-214

Aldisa tara Millen: 5(2):197-214

Aldisa trimaculata ‘Gosliner’ Millen and
Gosliner, 1985: 5(2):243-258

Aldisidae: 5(2):243-258

Alectryonella Sacco, 1897: 4(2):157-162

Alectryonella plicatula (Gmelin, 1791):
4(2):157-162

Aligena cokeri Dall, 1909: 1:91

Allogastropda: $1:1-22

Allogona profunda (Say, 1821): 1:97-98

Alloteuthis (Linneé, 1758): 4(2):217-227

Alvania abysicola (Forbes, 1850):
4(1):185-199 (passim)

Alvania (Alvania) isolata (Laseron, 1956):
4(2):232-233

Alvania auberiana (Orbigny, 1842):
4(2):185-199

Alvania punctura (Montagu, 1803):
4(1):185-199 (passim)

Amaea H. and A. Adams, 1853: $1:1-22

Amanda armata Macnae, 1954:
5(2):243-258

Amblema costata Rafinesque, 1820:
1:43-50; 6(1):19-37; S1:35-50

Amblema costata perplicata (Conrad,
1841): 6(1):19-37

Amblema costata plicata (Say, 1817):
6(1):19-37

Amblema peruviana: 6(1):19-37

Amblema plicata (Say, 1817): 1:29, 31-34,
43-50; 3(1):105; 4(1):25-37, 117;
5(2):165-171; 6(1):19-37, 49-54;
6(2):165-178

Amblema plicata plicata (Say, 1817):
1:51-60; 2:85-86; 3(1):47-53;
4(1):117-118; 6(1):19-37

Amblemidae Rafinesque, 1820: 4(1):117-188

Amblemini: 1:109-110

Ambloplites rupestris (Lacépede): 5(1):1-7

Amblychilepas Pilsbry, 1890: 2:21-34

Amete seftoni Berry, 1956: 3(1):63-82

Amianthus: 4(1):1-12

Ammonitellidae: 1:97

Ammonites: 2:79

Amnicola limosa (Say, 1817): 3(1):99;
5(1):9-19, 31-39, 73-84; 5(1):73-84
(passim)

Amnicola winkleyi Pilsbry, 1912:
4(1):101-102

Amoeba proteus: S1:79-83

Amphibola: $1:1-22

Amphibolidae: $1:1-22

Amphiroa: 4(2):185-199

Amphitretoidea Berry, 1920: 3(1):63-82

Amplirhagada \|redale, 1933: 1:98-99

Ampulla purpurea Roding, 1798: 2:57-61

Ampullariidae: 3(2):223-231

Amygdalum Muhlfeld, 1811: $1:23-34

Amygdalum politum (Verrill and Smith,
1880): S1:23-24

Anabaena: 4(1):81-88

Anabaena oscillarioides: S2:219-222

Anadara brasiliana (Lamarck, 1819): 4(1):111

Anadara (Cunearca) nux (Sowerby, 1857):
4(1):1-12

Anadara (Esmerarca) Olsson, 1961: 4(1):1-12

Anadara broughtonni (Schrenck, 1867):
4(1):111

Anadara granosa (Linné, 1758): 4(1):111

Anadara ovalis (Bruguiére, 1789): 4(1):111

Anadara transversa (Say, 1822): 4(1):111

Anaspidea: 4(1):109-110; 5(2):243-258;
$1:1-22

Anatina papyratia Say: 2:35-40

Ancipenser transmontanus Richardson:
$2:7-39

Ancistrobasis Dall, 1889: 1:92

Ancula Loven, 1846: 5(2):243-258

Ancula gibbosa (Risso, 1818): 5(2):185-196

Ancula pacifica MacFarland, 1905:
5(2):197-214

Anculosa: 4(1):25-37

Anculosa praerosa: 1:43-50

Ancylus drouetianus Bourguignat, 1853:
2:88-89

Ancylus fluviatilis Miller, 1776: 3(2):135-142,
151-168, 243-265, 269-272; 5(1):105-124

Ancylus gussonii Costa, 1829: 2:88-89

Anemonia sulcata Pennant: 5(2):185-196

Anguispira alternata (Say, 1816): 1:97-98;
3(1):27-32 (passim); 4(2):237; 6(1):16

AMER. MALAC. BULL. TAXONOMIC INDEX

Anguispira kochi (Pfeiffer, 1845): 1:97-98

Angutispira: $1:1-22

Anidolyta Gen. Nov., Willan, 1987:
§(2):216, 232-233

Anidolyta spongotheras Comb. Nov.,
Willan, 1987: 5(2):215-241

Anisodoris Bergh, 1898: 5(2):185-196

Anisodoris nobilis Macfarland, 1905:
5(2):197-214

Anisdoris prea Marcus and Marcus, 1967:
5(2):183-184

Ankistrodesmus: 4(1):81-88; S2:219-222

Ankylastrum capuloides: 5(1):65-72
(passim)

Ankylastrum fluviatile (Muller): 5(1):65-72
(passim)

Annelida: 3(2):213-221 (passim)

Anodonta sp.: 2:82; 4(1):13-19, 117-118;
$2:1-5; 6(2):179-188 (passim)

Anodonta anatina (Linné, 1758): 5(1):1-7

Anodonta cygnea Linne, 1758): 4(1):13-19;
5(1):41-48

Anodonta gibba Clessin, 1875: 5(1):91-99
(passim)

Anodonta grandis Say, 1829: 1:29, 43-50;
2:86; 3(1):93; 3(2):233-242; 5(1):91-99;
6(1):19-37; 6(2):165-178; S1:35-50

Anodonta grandis corpulenta Cooper, 1834:
1:51-60, 71-74; 5(1):31-39; 5(2):165-171;
6(1):19-37

Anodonta grandis gigantea Lea, 1838:
6(1):19-37

Anodonta grandis grandis Say, 1829: 1:51-60,
71-74; 2:85-86; 3(1):47-53, 105

Anodonta imbecilis Say, 1829: 1:51-60;
2:85-86; 3(1):47-53, 105; 4(1):21-23, 117;
4(2):231, 231-232; 6(1):19-37

Anodonta imbecilis henryiana (Lea, 1857):
2:86; 3(1):93

Anodonta implicata Say, 1829: 3(1):104-105;
4(1):13-19

Anodonta piscinalis Nilsson, 1822: 5(1):41-48

Anodonta subordiculata Say, 1831: 1:51-60,
71-74; 4(2):230-231; 6(1):19-37

Anodonta woodiana (Lea, 1834): 5(1):91-99

Anodontoides Baker, 1898: 4(1):117-118

Anodontoides ferussacianus (Lea, 1834):
3(1):93, 105; 5(2):165-171; 6(1):19-37

Anomalodesmata Dall, 1889: 4(1):111-112

Anomia Linneé, 1758: 4(2):157-162

Anomia simplex (Orbigny, 1842): 1:101-102;
2:41-50; S1:35-50

Anomiostrea Habe and Kosuge, 1966:
4(2):157-162

Anomiostrea coralliophila Habe, 1975:
4(2):157-162

Anthobranchia: 5(2):215-241

Anthopleura elegantissima (Brandt):
5(2):287-292

Antiopella barbarensis (Cooper, 1863):
5(2):287-292

Antiplanes (Ractiplanes) willetti Berry, 1953:
3(1):63-82

Antiplanes macfarlandi Berry, 1947:
3(1):63-82

Antonietta luteorufa Schmekel: 5(2):197-214

Aphanistylus Fischer, 1884: 2:1-20

Aphelodoris brunnea Bergh, 1907:
5(2):243-258

Aphrodita: 1:90-91

Aplacophora von Ihering, 1876:
3(1):93-94; 4(1):107; 5(2):281-286;
$1:23-24; $1:35-50

Aplocinotus grunniens Rafinesque):
S2:7-39, 89-94

Aplysia sp.: 2:78; 5(2):185-196; S1:1-22

Aplysia brasiliana Rang, 1828: 2:78

Aplysia californica Cooper, 1863: 2:78

Aplysia dactylomela Rang, 1825:
5(2):243-258

Aplysia juliana Quoy and Gaimard, 1832:
5(2):197-214, 243-258

Apiysia oculifera Adams and Reeve,
1850: 5(2):243-258

Aplysia parvula Guilding?: 5(2):185-196

Aplysia parvula Morch, 1863:
5(2):243-258

Aplysia punctata: 4(2):205-216 (passim)

Aplysiidae Rafinesque, 1815: 5(2):243-258;
$1:1-22

Aplysiomorpha: $1:1-22

Aplysiopsis sinusmensalis (Macnae,
1954): 5(2):243-258

Aplysiopsis smithi (Marcus): 5(2):197-214

Aplysiopsis zebra Clark: 5(2):259-280

Arca noae Linné, 1758: S1:59-78

Arcacea Lamarck, 1809: 2:41-50

Archaeogastropoda Thiele, 1925:
$1:23-24

Archiconchifera: 6(1):57-68

Archidoris Bergh, 1878: 5(2):185-196

Archidoris britannica (Leach, 1852):
4(2):205-216 (passim)

Archidoris monteryensis (Cooper, 1862):
4(2):205-216 (passim); 5(2):185-196

Archidoris odhneri (MacFarland, 1966):
5(2):197-214

Archidoris pseudoargus (Rapp, 1827):
4(1):103-104; 4(2):205-216 (passim),
232; 5(2):185-196, 197-214

Archiplacophora: 6(1):57-68

Architectonica (Architectonica) Roding,
1798): 4(1):108-109

Architectonicacea: $1:1-22

Architectonicidae Gray, 1850: 4(2):236;
$1:1-22

Architeuthoidea Berry, 1920: 3(1):63-82

Arcidens confragosus (Say, 1829):
1:51-60; 5(2):165-171; 6(1):19-37

Arctica islandica (Linné, 1767): S1:59-78;
$3:51-57

Arcticacea Newton, 1891: 3(1):103

Arcuatula ‘Jousseaume’ Lamy, 1919:
5(2):159-164

Arenicola: 2:96

Argonauta Linné, 1758: 4(2):217-227

: 1983 - 1988 225

Argonauta argo Linne, 1758: 5(2):303-306

Argonautoidea Berry, 1920: 3(1):63-82

Argopecten arquisulcatus: 4(2):241-242

Argopecten gibbus (Linné, 1758): 2:41-50

Argopecten irradians (Lamarck, 1819):
$1:59-78

Arianta arbustorum: 1:103

Ariolimax colmbianus (Gould, 1851):
$1:35-50

Arion ater Linne, 1758): 1:110; 3(1):27-32
(passim); 6(1):16

Arion ater rufus (Linné, 1758): 6(1):16

Arion circumscriptus Johnston, 1828:
1:110; 6(1):16

Arion distinctus Mabille: 1:110; 6(1):16

Arion hortensis Férussac, 1819: 6(1):16

Arion intermedius (Normand, 1852): 1:110;
6(1):16

Arion lusitanicus Mabille: 6(1):16

Arion owenii Férussac, 1819: 6(1):16

Arion silvaticus Lohmander: 1:110; 6(1):16

Arion subfuscus (Draparnaud, 1805): 1:24
(passim), 1:110; 6(1):16

Arionidae Gray, 1840 : S1:35-50

Armina Rafinesque, 1814: 5(2):185-196

Armina californica (Cooper, 1862):
5(2):197-214

Armina gilchristi (Bergh, 1907): 5(2):243-258

Armina maculata Rafinesque, 1814:
5(2):197-214

Armina tigrina Rafinesque, 1814:
4(2):205-216 (passim)

Arminacea Rafinesque, 1814: 5(2):215-241

Arminidae Rafinesque, 1814: 5(2):243-258

Artachaea Bergh, 1882: 5(2):243-258

Arthritica hulmei Ponder, 1965: 1:90-91

Arthropoda: 3(2):213-221 (passim)

Ascobulla fischeri (Adams and Angas,
1864): 5(2):243-258

Ascobulla ulla (Marcus and Marcus):
5(2):259-280

Ascoglossa Bergh, 1877: $1:1-22

Ascophyllum: 1:92

Ascoteuthis Berry, 1920 : 3(1):63-82

Ashmunella chiricahuna Dall, 1895: 1:98;
2:98

Ashmunella lenticula Gregg, 1953: 1:106

Ashmunella levettei (Bland, 1880): 1:21-26

Ashmunella proxima albicaudata Pilsbry
and Ferriss, 1910: 1:106

Ashmunella varicifera (Ancey, 1901):
1:21-26

Asparagopsis taxiformis: 5(2):185-196

Aspidodiadema hawaiiensis: 2:83

Assiminea californica (Tryon, 1865):
4(1):185-199 (passim)

Assiminea infima Berry, 1947: 3(1):63-82

Assimineidae H. and A. Adams, 1856:
3(2):223-231

Astarte castanea (Say, 1822): 5(1):21-30
(passim); S1:59-78

Astrea (Pomaulax) petrohauma Berry,
1940: 3(1):63-82

226 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Astraea guadalupeand Berry, 1940:
3(1):63-82

Astraea rugosa (Linné, 1758): 5(2):303-306

Asterias amurensis: 2:94

Asterias forbesi (Desor): S3:59-70

Asterionella: S2:167-178

Asteronotidae: 5(2):243-258

Ataagena: 5(2):185-196

Atagema gibba Pruvot-Fol, 1951:
5(2):243-258

Atagema rugosa Pruvot-Fol, 1951:
5(2):243-258

Atrina seminuda (Lamarck, 1819): 2:97

Atyidae Thiele, 1926: 4(2):233; S1:1-22

Atys Montfort, 1810: 5(2):185-196

Atys cylindrica (Helbling, 1779):
5(2):243-258

Aufwuchs: 3(2):169-177, 243-265

Australorbis glabratus (Say, 1818):
3(2):213-221

Austrocochlea constricta Fisher: 6(1):17

Austrodoris macmurdensis Odhner, 1934:
4(2):205-216 (passim)

Austrophon Dall, 1902: 3(1):11-26

Austrossia Berry, 1918: 3(1):63-82

Avicennia: 4(1):112

Avrainvillea nigricans Decaisne: 5(2):259-280

Axinulus Verrill and Bush, 1898: 2:96

Axinulus brevis: 2:96

Aythya affinis (Eyton): S3:59-70

Aythya marila (Linné, 1758): S3:59-70

Babaina: 5(2):197-214

Bacillariophycaea: S2:167-178

Baeolidida palythoae Gosliner, 1985:
5(2):243-258

Balanus amphitrite amphitrite Darwin:
$1:111-116

Balanus concavus Bronn, 1831: 4(1):39-42

Balanus finchii Lea, 1833: 4(1):39-42

Balanus improvisus: S2:133-142 —

Balanus proteus Conrad, 1834: 4(1):39-42

Balcis (Balcis) clavella Berry, 1954:

3(1):63-82

Balcis (Balcis) tersa Berry, 1954:
3(1):63-82

Balcis (Vitreolina) ebriconus Berry, 1954:
3(1):63-82

Balcis (Vitreolina) incallida Berry, 1954:
3(1):63-82

Balcis (Vitreolina) obstipa Berry, 1954:
3(1):63-82

Balcis (Vitreolina) titubans Berry, 1954:
3(1):63-82

Bankia Gray, 1842: 3(1):85-88

Bankia gouldi Bartsch, 1908: 4(1):89-99;
$1:101-109

Bankivia Menke, 1830: 3(1):95

Barbatia (Acar) rostae Berry, 1954:
3(1):63-82

Barleeia sp.: 4(2):232-233

Basiliochiton Berry, 1918: 3(1):63-82

Basiliochiton lobium Berry, 1925:
3(1):63-82

Basommatophora Keferstein, 1864: S1:1-22

Bathybemix bairdii (Dall, 1889): 6(1):9-17

Bathyberthella Willan, 1983: 5(2):215-241

Bathyberthella antarctica Willan and
Bertsch, 1987: 5(2): 215-241

Bathyberthella zelandiae Willan, 1983:
5(2):215-241

Bathydorididae: 5(2):243-258

Bathypolypus arcticus (Prosch, 1849):
4(2):217-227

Bathyteuthis Hoyle, 1885: 3(1):55, 56
(passim)

Bathyteuthis berryi Roper, 1954: 3(1):55,
56 (passim)

Batillaria Benson, 1842: 2:1-20

Batillaria minima (Gmelin, 1791): 2:1-20

Batillaria zonalis (Bruguiere, 1792): 2:1-20

Batillariinae Thiele, 1929: 2:1-20

Batissa (Cyrenobatissa) subsulcata
Clessin, 1878: 5(1):91-99

Bellamya capillata: 4(1):107

Bellamya jeffreysi: 4(1):107

Bellamya unicolor: 4(1):107

Benthoteuthidae Berry, 1912: 3(1):63-82

Benthoteuthis Verrill, 1885: 3(1):56
(passim)

Bernardina bakeri Dall, 1910: 3(1):103

Bernardina margarita (Carpenter, 1857):
3(1):103

Bernardinidae Keen, 1963: 3(1):103

Berryteuthis anonychus (Pearcy and Voss,
1963): 2:89; 4(2):241

Berryteuthis magister (Berry, 1913): 2:89

Berthelinia Crosse, 1875: S1:1-22

Berthelinia caribbea Edmunds, 1963:
5(2):197-214, 259-280

Berthelinia limax Kawaguti and Baba:
5(2):197-214

Berthelinia schlumbergeri Dautzenberg,
1895: 5(2):243-258

Berthella Blainville, 1825: 5(2):215-241;
$1:1-22

Berthella americana (Verrill): 5(2):215-241

Berthella californica (Dall, 1900):
5(2):197-214

Berthella martensi (Pilsbry, 1896):
5(2):215-241

Berthella medietas Burn: 5(2):215-241

Berthella ornata (Cheeseman):
5(2):215-241

Berthella pellucida (Pease): 5(2):215-241

Berthella plumula (Montagu, 1803):
5(2):215-241, 243-258

Berthella porosa Blainville, 1825:
5(2):215-241

Berthella stellata (Risso): 5(2):185-196

Berthella tupala Marcus, 1957:
5(2):243-258

Berthellina Gardiner, 1936: 5(2):215-241;
$1:1-22

Berthellina citrina (Ruppell and Leuckart,
1828): 5(2):197-214, 215-241,
243-258

Berthellina engeli Gardiner, 1936:
§(2):215-241

Berthellinae Burn, 1962: 5(2):215-241

Berthellini: 5(2):215-241

Berthellinops Burn, 1962: 5(2):215-241

(Bessomia) Berry, 1959: 3(1):63-82

Bimeria: 5(2):197-214

Biomphalaria alexandria (Ehrenberg):
6(1):17

Biomphalaria alexandrina (Bourguignat,
1883): 1:67-70, 107

Biomphalaria boissyi: 1:67-70

Biomphalaria choanomphala (Martens,
1879): 5(1):85-90

Biomphalaria glabrata (Say, 1818): 1:67-70,
96-97, 106, 106-107, 107; 3(1):89-90;
3(2):213-221; 4(1):120; 5(1):65-72;
105-124 (passim); 6(1)17; S1:25-50,
79-83

Biomphalaria havanensis (Pfeiffer): 6(1):17

Biomphalaria pfeifferi (Krauss, 1848):
5(1):65-72, 85-90; 105-124 (passim)

Biomphalaria sudanica (Martens, 1870):
5(1):85-90

Biomphalaria stanleyi (Smith, 1888):
5(1):85-90

Biomphalaria straminea (Dunker): 1:67-70,
106-107; 6(1):17

Biomphalaria tenagophila: 1:67-70

Bithynia Leach, 1818: 3(2):135-142
(passim), 269-272

Bithynia tentaculata (Linné, 1758):
3(2):179-186

Bithyniidae Walker, 1927: 3(2):223-231

Bittium Gray, 1847: 2:1-20

Bittium alternatum (Say, 1822): S1:85-91

Bittium varium Pfeiffer, 1840: 4(2):185-199

Bivalvia, Unspecified: 3(1):93, 93-94;
4(1):102-103, 111-112; S2:69-81

Bivetiella Wenz, 1943: 2:57-61

Blauneria Shuttleworth, 1854: $1:1-22

Boccardia ligerica: S2:7-39

Bonsia nakaza Gosliner, 1981:
5(2):243-258

Boonea Robertson: S1:1-22; S3:59-70

Boonea impressa (Say, 1821): 3(1):97;
$3:59-70

Booneostrea: 4(2):157-162

Booneostrea cucullina (Deshayes, 1836):
4(2):157-162

Boreotrophon aculeatus (Watson, 1882):
3(1):11-26

Boreotrophon alborostratus Taki, 1938:
3(1):11-26

Boreotrophon lacunellus (Dall, 1889):
3(1):11-26

Boreotrophon truncatus (Strom, 1768):
3(1):11-26

Bornella anguilla Johnson, 1983:
5(2):243-258

Bornella stellifer (Adams and Reeve’ A.
Adams, 1848): 5(2):243-258

Bornellidae: 5(2):243-258

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 2ef

Bosellia mimetica Trinchese: 5(2):197-214,
259-280

Bosellidae: 5(2):259-280

Botula cylista Berry, 1959: 3(1):63-82

Boveria teredinidi: S1:101-109

Boveria zeukevitchi Levinson: S1:101-109

Brachidontes exustus (Linné, 1758):
4(2):233-234

Bradybaenidae: 2:97

Bradybaena similaris Ferussac: 2:97; 6(1):16

Bradybaena (Acusta) despecta sieboldiana:
2:97

Brechites penis (Linné, 1758): 5(1):21-30
(passim)

Brondelia Bourguignat, 1862: 2:89-90

Brondelia drouetiana (Bourguignat, 1853):
2:89-90

Brondelia gibbosa Bourguignat, 1862:
2:89-90

Bryopsis: 5(2):259-280

Bryopsis plumosa (Hudson) Agardh:
5(2):259-280

Buccinacea Rafinesque, 1815: 3(1):11-26

Buccinanops: 3(1):101-102

Buccinum Linné, 1758: $1:35-50

Buccinum evulsum Solander, 1766:
2:57-61

Buccinum piscatorium Gmelin, 1791:
2:57-61

Buccinum pyrozonias Gmelin, 1791:
2:57-61

Buccinum scalare Gmelin, 1791: 2:57-61

Buccinum undatum Linne, 1758:
3(2):223-231; 4(1):185-199 (passim)

Buchanaania Gistel, 1848: 2:21-34

Buchanania Lesson, 1830: 2:21-34

Buchanania onchidioides Lesson, 1830:
2:21-34

Bulimulidae: 1:97; 3(1):8 (passim);
4(1):113-114

Bulinus cernicus: 1:107

Bulinus forskali (Ehrenberg, 1831)-Group:
1:107

Bulinus jousseaumei (Dautzenberg, 1890):
5(1):65-72

Bulinus natalensis ‘Krauss’ Kuster,
1841-1843: 1:107, 106-107

Bulinus tropicus (Krauss, 1848): 1:96,
106-107

Bulinus truncatus (Audouin): 1:106-107;
5(1):85-90

Bulla Linneé, 1758: 5(2):185-196; $1:1-22

Bulla ampulla (Linné, 1758): 5(2):243-258

Bulla membranacea Montagu, 1815:
5(2):215-241

Bulla plumula Montagu, 1803: 5(2):215-241

Bullia: 3(1):101-102

Bullidae Rafinesque, 1815: 4(2):233;
5(2):243-258; $1:1-22

Bullina Férussac, 1822: 5(2):185-196;
$1:1-22

Bullina lineata (Gray, 1825): 5(2):243-258

Bullinidae: 5(2):243-258

Bullomorpha: S1:1-22

Bursa californica sonorana Berry, 1940:
3(1):63-82

Bursatella Blainville, 1817: 5(2):185-196

Bursatella leachii africana (Engel, 1927):
5(2):243-258

Bursatella leachii leachii (Blainville, 1817):
5(2):243-258

Busycon sp.: 4(1):25-37, 185-199 (passim);
$1:35-50; S3:59-70

Busycon canaliculatum (Linné, 1758):
3(1):27-32 (passim), 102; S3:59-70

Busycon carica (Gmelin, 1791): 3(1):27-32
(passim), 102; S3:59-70

Busycon contrarium (Conrad, 1840):
3(1):102; 4(1):110

Busycon spiratum (Lamarck, 1816):
3(1):102

Bythinia tentaculata (Linné, 1758):
5(1):65-72 (passim); S2:1-5 (passim)

Cadlina Bergh, 1879: 5(2):243-258

Cadlina laevis (Linné, 1767): 4(1):103-104;
4(2):205-216 (passim); 5(2):197-214

Cadlina modesta MacFarland, 1966:
5(2):197-214

Caecum Fleming, 1813: 5(2):281-286

Caecum nitidum Stimpson, 1851:
4(1):185-199

Caecum septimentum deFolin, 1867:
4(2):232-233

Caelatura Conrad, 1853: 4(1):107

Calcitrapessa Berry, 1959: 3(1):63-82

Caliphyliidae Tiberi, 1880: 5(2):243-258

Caliphylla mediterranea Costa, 1867:
5(2):197-214, 259-280

Caliphyllidae Tiberi, 1880: 5(2):259-280

Callinectes sapidus (Rathburn): S3:51
(passim), 59-70

Calliopaea bellula (Orbigny, 1837):
5(2):197-214

Calliostoma apicinum Dall, 1881: 2:84

Calliostoma grantianum Berry, 1940:
3(1):63-82

Calliostoma hannibali Hertlein and Jor-
dan, 1927: 4(1):1-12

Calliostoma pulchrum (C. B. Adams,
1850): 2:84

Calliostoma roseolum Dall, 1881: 2:84

Calliostoma velieli Pilsbry, 1900: 2:84

Calliostoma zizyphinum (Linnée, 1758:
4(1):185-199 (passim)

Callistochiton ‘Carpenter’ Dall, 1878:
6(1):115-130

Callistochiton adenensis Smith, 1891:
6(1):115-130

Callistochiton barnardi Smythe, 1982:
6(1):115-130

Callistochiton decoratus ferminicus Berry,
1922: 3(1):63-82

Callistochiton finschi Thiele, 1910:
6(1):115-130

Callistochiton heterodon savignyi Pilsbry,
1893: 6(1):115-130

Callistochiton palmulatus ‘Carpenter’ Dall,
1879: 6(1):115-130

Callistochitoninae Berry, 1922: 3(1):63-82

Callistoplacinae Pilsbry, 1893: 6(1):115-130

Calliteuthis (Meleagroteuthis) heteropsis
Berry, 1913: 3(1):63-82

Calliteuthis miranda Berry, 1918:
3(1):63-82

Callochitonidae Plate, 1899: 6(1):141-151

Calma glaucoides (Alder and Hancock,
1854): 5(2):197-214

Calmella carolinii Verany: 5(2):185-196,
197-214

Calocochlea: 3(1):98-99

Calocochlea caillaudi (Deshayes):
3(1):98-99

Caloria Trinchese, 1888: 5(2):243-258

Caloria indica (Bergh, 1896): 5(2):243-258

Calotrophon ostrearum (Conrad, 1846):
4(1):185-199 (passim)

Calyptogena Dall, 1891: S1:23-24

Calyptogena magnifica Boss and Turner,
1980: 1:101; 4(1):49-54; $1:23-34

Calyptogena ponderosa Boss, 1968:
$1:23-24

Calyptraeide: 3(1):85-88; 4(2):173-183;
$1:1-22

Calyptraea Lamarck, 1799: 4(1):1-12

Calyptraea chinensis (Linné, 1758):
3(2):179-186 (passim)

Calyptraea conica Broderip, 1834:
4(2):173-183

Calyptraea mamillaris Broderip:
4(2):173-183

Calyptraea novazelandiae: 4(2):173-183

Camaenidae: 3(1):8 (passim)

Cambarus bartonii: S2:89-94, 211-218

Campanile: $1:1-22

Campanilidae: $1:1-22

Campeloma sp.: 1:43-50; 4(1):25-37

Campeloma crassula (Rafinesque, 1819):
4(1):25-37

Campeloma decisum (Say, 1816):
4(1):25-37; 5(1):9-19, 31-39, 73-84,
101-104; 6(1):17; 6(2):165-178

Campeloma exile (Anthony, 1860):
4(1):25-37

Campeloma geniculum (Conrad, 1834):
3(1):99; 4(1):25-37; 6(1):17

Campeloma parthenum Vail, 1979:
3(1):99; 6(1):17

Campeloma ponderosum (Cooper, 1834):
4(1):25-37

Campeloma rufum (Haldeman, 1841):
4(1):25-37

Campostoma anomalum (Rafinesque):
5(1):1-7

Cancellaria Lamarck, 1799: 2:57-61

Cancellaria (Bivetiella) cancellata (Linne,
1767): 2:57-61

Cancellaria cancellaria (Linné, 1758):
2:57-61

Cancellaria costata Sowerby, 1821: 2:57-61

228 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Cancellaria costata Sowerby, 1833: 2:57-61

Cancellaria lamellosa Hinds, 1843: 2:57-61

Cancellaria nassa (Gmelin, 1791): 2:57-61

Cancellaria nodulosa Lamarck, 1822:
2:57-61

Cancellaria (Pyruclia) diadela: 2:84-85

Cancellaria reticulata (Linné, 1767): 2:57-61

Cancellaria scalarina Lamarck, 1822:
2:57-61

Cancellaria similis Sowerby, 1833: 2:57-61

Cancellaria (Solatia) piscatoria (Gmelin,
1791): 2:57-61

Cancellaria reticulata (Linné, 1767): 4(1):113

Cancellaria trigonostoma (Lamarck,
1822): 2:57-61

Cancellariidae Forbes and Handley, 1853:
2:57-61

Cantharus multangulus (Philippi, 1848):
4(1):185-199 (passim)

Cantharus rehderi Berry, 1962: 3(1):63-82

Cantharus shaskyi Berry, 1959: 3(1):63-82

Cantharus triplicatus Roding, 1798: 2:57-61

Capitella capitata: S2:203-209

Capulidae Fleming, 1822: S1:35-50

Capulis ungaris (Linne, 1767):
3(2):179-186 (passim)

Caracolus: 3(1):8 (passim)

Carcinus maenas: 4(1):108

Cardiomya planetica (Dall, 1908): 1:13

Cardita (Cardites) Link, 1807: 4(1):1-12

Cardium 6(2):165-178 (passim)

Cardium edule Linné, 1758: 3(1):33-40

Caretta caretta: 3(1):93

Caruncula moesta (Lea, 1841): 1:43-50

Caruncula moesta cylindrella (Lea, 1868):
1:43-50

Caruncula parva (Barnes, 1823): 3(1):105

Carunculina glans (Lea, 1831): 6(1):19-37

Carunculina lividus (Simpson, 1900):
1:43-50

Carunculina moesta (Lea, 1841): 6(1):19-37

Carunculina moesta cylindrella (Lea,
1868): 6(1):19-37

Carunculina parva (Barnes, 1823):
6(1):19-37

Carunculina texasensis (Lea, 1857):
3(2):233-242

Carychium (Miller, 1774): $1:1-22

Casella obsoleta (Ruippell and Leuckart,
1831): 5(2):197-214

Cassiopea frondoza Fuwkes: 5(2):185-196

Cassiopea xamachana Bigelow:
5(2):185-196

Cassis tuberosa (Linné, 1758): $1:35-50

Catostomus commersoni: S2:69-81

Catriona casha Gosliner and Griffiths,
1981: 5(2):243-258

Catriona gymnota (Couthouy, 1838):
5(2):185-196, 197-214, 287-292

Catriona maua Marcus and Marcus,
1960: 5(2):183-184, 197-214

Caudofoveata: 4(1):107; 6(1):57-68

Caulerpa: 5(2):185-196

Caulerpa mexicana (Sonder) Kitzing:
5(2):259-280

Caulerpa okamurai (Webber-Van Basse):
5(2):197-214

Caulerpa paspaloides (Bory) Greville:
5(2):259-280

Caulerpa racemosa (Forsskal) Agardh:
5(2):259-280

Caulerpa sertulariodes (Webber-van
Bosse) Borgesen: 5(2):259-280

Caulerpa verticilliata (Agardh):
5(2):197-214, 259-280

Cellana: 4(1):115

Cepaea sp.: 1:103; 6(1):9-17

Cepaea hortensis (Miiller, 1774): 1:97-98,
103; 6(1):16

Cepaea nemoralis (Linné, 1758): 1:97-98,
103, 107-108; 3(1):1-10; 5(2):105-124;
6(1):16

Cepaea nemoralis nemoralis (Linne,
1758): 1:107-108

Capaea sylvatica Draparnaud: 6(1):16

Cepaea vindobonensis: 1:107-108

Cephalaspidea P. Fischer, 1883: 4(2):233;
5(2):243-258; S$1:1-22

Cephalopoda, Unspecified: 2:89, 2:90-91;
6(1):57-68 (passim)

Ceratium hirundinella: S2:167-178

Ceratophyllidia africana Eliot, 1903:
5(2):243-258

Ceratosoma Hermannsen, 1846:
5(2):243-258

Ceratosoma cornigerum A. Adams and
Reeve, 1820: 5(2):243-258

Ceratozona squalida (C. B. Adams, 1845):
6(1):79-114

Cerberilla Bergh, 1873: 5(2):185-196

Cerion Roding, 1798: 6(1):9-17

Cerion bendalli Pilsbry and Vanatta:
6(1):16

Cerion incanum (Binney, 1851) : 6(1):16

Cerithdeopsilla: 2:1-20

Cerithiacea Fleming, 1822: 2:1-20

Cerithidea s.s.: 2:1-20

Cerithidea Swainson, 1840: 2:1-20; 3(1):59
(passim)

Cerithidea alata: 2:1-20

Cerithidea californica (Haldeman, 1840):
2:1-20; 4(2):165-172

Cerithidea (Cerithideopsilla) Thiele, 1929:
2:1-20

Cerithidea (Cerithideopsis) Thiele, 1929:
2:1-20

Cerithidea Charbonieri (sic) Petit de la
Saussaya, 1851: 1:1-20

Cerithidea Charbonniere (sic) Petit de la
Saussaya, 1851: 2:1-20

Cerithidea cingulata (Gmelin, 1807):
2:1-20

Cerithidea costata (da Costa, 1778):
2:1-20

Cerithidea decollata (Linné, 1767): 2:1-20

Cerithidea djadjariensis: 2:1-20

Cerithidea fluviatilis (Potiez and Michaud,
1838): 2:1-20

Cerithidea iostoma (Pfeiffer, 1829): 2:1-20

Cerithidea kieneri: 2:1-20

Cerithidea largillierti Philippi, 1849: 2:1-20

Cerithidea lutosum (Menke): 2:1-20

Cerithidea microptera (Kiener): 2:1-20

Cerithidea modulus Say: 2:1-20

Cerithidea montagnei (Orbigny, 1841):
2:1-20

Cerithidea muscarum: 2:1-20

Cerithidea obtusa (Lamarck, 1822): 2:1-20

Cerithidea pliculosa (Menke, 1822):
2:1-20

Cerithidea quadrata Sowerby, 1855:
2:1-20

Cerithidea reevianum C. B. Adams: 2:1-20

Cerithidea rhizophorarum: 2:1-20

Cerithidea sacrata hyporhyssa Berry,
1906: 3(1):63-82

Cerithidea scalariformis (Say, 1825):
2:1-20; 4(1):111; 4(2):234

Cerithideopsis Thiele, 1929: 2:1-20

Cerithiidae Fleming, 1828: 2:1-20;
3(2):223-231; 4(2):235

Cerithiopsacea: $1:1-22

Cerithiopsidae: S1:1-22

Cerithium Bruguiére, 1789: 4(1):1-12;
6(1):9-17

Cerithium caeruleum Sowerby, 1855:
6(1):17

Cerithium ebininum: 2:1-20

Cerithium nodulosum Bruguiere, 1789:
2:1-20

Cerithium obtusa (Lamarck, 1822): 2:1-20

Cerithium placidum Gould, 1849:
4(2):232-233

Cerithium rupestre (Risso): 6(1)17

Cerithium scabridum Philippi: 6(1):17

Chaetodermomorpha ‘Pelseneer’ Lank-
ester, 1906: 6(1):57-68

Chaetoderma: 6(1):57-68

Chaetogaster limnaei limnaei:
3(2):151-168; S2:7-39, 89-94

Chaetomorpha: 5(2):259-280

Chaetopleura angulata (Spengler, 1797):
6(1):115-130

Chaetopleura apiculata (Say, 1834):
4(1):107-108; 6(1):69-78

Chaetopleura lurida (Sowerby, 1832):
6(1):141-151

Chaetopleura peruviani Lamarck:
6(1):141-151

Chaetopleura (Pallochiton) euryplax Berry,
1945: 3(1):63-82

Chaetopleuridae Plate, 1899: 6(1):141-151

(Chamaearionta) Berry, 1930: 3(1):63-82

Charonia tritonis (Linné, 1758): 2:84

Charopidae: 2:97

Chelidonura A. Adams, 1850: 5(2):197-214;
$1:1-22

Chelidoneura fulvipunctata Baba, 1938:
5(2):243-258

AMER. MALAC. BULL. TAXONOMIC INDEX

Chelidoneura hirudinina (Quoy and
Gaimard, 1824): 5(2):243-258

Chicoreus palmarosae Lamarck, 1822:
3(1):11-26

Chicoreus virgineus (Roding, 1798):
4(1):109-110

Chilina: $1:1-22

Chilinidae: $1:1-22

Chilomonas: S1:79-83

Chione Mihlfeld, 1811: 4(1):1-12

Chione cancellata (Linné, 1758): 2:41-50;
4(1):111

Chione (Chione) richthofeni Hertlein and
Jordan, 1927: 4(1):1-12

Chione (Chionopsis) Olsson, 1932:
4(1):1-12

Chiroteuthis famelica Berry, 1909:
3(1):63-82

Chiroteuthoidea Berry, 1920: 3(1):63-82

Chiroteuthoides Berry, 1920: 3(1):63-82

Chiroteuthoides hastula Berry, 1920:
3(1):63-82

Chiton Linné, 1758: 2:21 (passim);
4(1):114-115; 6(1):115-130

Chiton affinis |ssel, 1869: 6(1):115-130

Chiton astringer Reeve, 1847: 1:91;
6(1):79-114

Chiton chilensis Frembly, 1827: 6(1):115-130

Chiton confossus Gould, 1846: 6(1):115-130

Chiton elegans Frembly, 1827: 6(1):115-130

Chiton fascicularis Linne, 1767: 6(1):115-130

Chiton fosteri Bullock, 1972: 6(1):115-130

Chiton huluensis (Smith, 1903): 6(1):115-130

Chiton iatricus Winckworth, 1930:
6(1):115-130

Chiton iatricus winckworthi Kaas, 1954:
6(1):115-130

Chiton janeirensis Gray, 1828: 6(1):115-130

Chiton lamellosus Quoy and Gaimard,
1835: 6(1):115-130

Chiton lamyi Dupuis, 1917: 6(1):115-130

Chiton lamyi reticulatus Dupuis, 1918:
6(1):115-130

Chiton luzonicus Sowerby, 1842:
6(1):115-130

Chiton mertensii Middendorff, 1847:
6(1):115-130

Chiton olivaceus Spengler, 1797:
6(1):131-139, 141-151, 153-159

Chiton olivaceus affinis \ssel, 1869:
6(1):115-130

Chiton peregrinus Thiele, 1910: 6(1):115-130

Chiton polii (Philippi): 6(1):57-68

Chiton punctatus Linné, 1758: 6(1):115-130

Chiton salihafui Bullock, 1972: 6(1):115-130

Chiton sueziensis Reeve, 1847:
6(1):115-130

Chiton spiculosus Reeve, 1847: 1:91;
6(1):79-114

Chiton spinosus Bruguiére, 1792:
6(1):115-130

Chiton squamosus Linné, 1764: 6(1):79-114

Chiton strigatus Sowerby, 1840: 6(1):79-114

Chiton testudo Spengler, 1797: 6(1):115-130

Chiton textilis Gray, 1828: 6(1):115-130

Chiton tuberculatus Linné, 1758:
6(1):115-130

Chiton wallacei Winckworth, 1927:
6(1):115-130

Chiton (Acanthopleura) haddoni (Winck-
worth, 1927): 6(1):115-130

Chiton (Callistochiton) adenensis Smith,
1891: 6(1):115-130

Chiton (Chiton) fosteri Bullock, 1972:
6(1):115-130

Chiton (Chiton) peregrinus Thiele, 1910:
6(1):115-130

Chiton (Clathropleura) peregrinus Thiele,
1910: 6(1):115-130

Chiton (Ischnochiton) yerburyi Smith,
1891: 6(1):115-130

Chiton (Rhyssoplax) affinis Issel, 1869:
6(1):115-130

Chiton (Rhyssoplax) olivaceus Spengler,
1797: 6(1):115-130

Chiton latus Guilding, 1829: 6(1):79-114

Chitonidae Rafinesque, 1815: 4(1):114-115;
6(1):115-130, 141-151

Chitoninae Linné, 1758: 6(1):115-130

Chlamydomonas: 4(1):81-88

Chlamys islandica (Miller, 1776): S1:35-50

Chlamys opercularis (Linné, 1758): 1:13
(passim)

Chlorella: 4(1):81-88; S2:143-150, 167-178

Chlorella vulgaris: 3(2):179-186;
$2:219-222

Chlorophyceae: S2:167-178

Chondrocidaris gigantea: 2:83

Choneplax Dall, 1882: 6(1):79-114

Choneplax lata (Guilding, 1829):
6(1):79-114

Choromytilus palliopunctatus (Carpenter,
1897): 4(1):1-12

Chromodorididae: 5(2):243-258

Chromodoris Alder and Hancock, 1855:
5(2):197-214, 287-292

Chromodoris africana Eliot, 1904:
5(2):243-258

Chromodoris albopunctatus (Garrett,
1897): 5(2):197-214, 287-292

Chromodoris alderi Collingwood, 1881:
5(2):243-258

Chromodoris annulata Eliot, 1904:
5(2):243-258

Chromodoris aspersa (Gould, 1852):
5(2):243-258

Chromodoris diardii (Kelaart, 1857):
5(2):185-196

Chromodoris elegantula Philippi, 1844:
5(2):185-196

Chromodoris geometrica (Risbec, 1928):
5(2):243-258

Chromodoris hamiltoni Rudman, 1977:
5(2):243-258

Chromodoris inopinata Bergh, 1905:
5(2):243-258

: 1983 - 1988 220

Chromodoris inornata Pease, 1871:
4(1):109-110; 5(2):197-214

Chromodoris krohni (Verany, 1846):
5(2):185-196, 197-214

Chromodoris loringi (Angas, 1864):
5(2):197-214

Chromodoris luteopunctata (Gantés,
1862): 5(2):197-214

Chromodoris marginata (Pease, 1860:
5(2):243-258

Chromodoris quadricolor Ruppell and
Leuckart, 1831: 4(1):109-110

Chromodoris reticulata (Pease, 1860):
5(2):185-196

Chromodoris tryoni (Garrett, 1873):
5(2):197-214

Chromodoris vicina Eliot, 1904:
5(2):243-258

Chromodoris sp.: 5(2):243-258

Chrysallida Carpenter, 1857: S1:1-22

Chrysaora quinquecirrha (Desor) S3:59-70

Chrysophyceae: S2:167-178

Cimora coneja Marcus, 1961: 5(2):287-292

Cincinnatia cincinnatiensis (Anthony,
1840): 5(1):31-39, 105-124 (passim)

Cincinnatia winkleyi (Pilsbry, 1912):
4(1):101-102

Cingula Fleming, 1828: 4(1):185-199
(passim)

Cionella lubrica (Muller): 3(1):27-32;
$1:35-50

Cipangopaludina chinensis (Gray, 1834):
5(1):9-19

Cirostrema pentedesmium Berry, 1963:
3(1):63-82

Cirroteuthis macrope Berry, 1911:
3(1):63-82

Cirroteuthoidea Berry, 1920: 3(1):63-82

Cistopus indicus (Orbigny): 6(2):207-211

Cladobranchia: 5(2):215-241

Cladophora Gary, 1840: 5(2):259-280

Cladophora gracilis (‘Griffiths’ Harvey)
Kuitzing: 5(2):259-280 (passim)

Cladophora prolifera (Roth) Kiitzing:
5(2):259-280

Cladophoropsis: 5(2):259-280

Clathrina coriacea (Montagu): 5(2):185-196

Clathurella (Glyphostoma) tridesma Berry,
1941: 3(1):63-82

Clavagella australis Sowerby, 1829:
$1:35-50

Clavagellidae Orbigny, 1843: S1:35-50

Clavus (Crassispira) zizyphus Berry, 1940:
3(1):63-82

Cleanthus Gray, 1847: 5(2):215-241

Cleidothaeridae Hedley, 1918: S1:35-50

Cliona celata Grant: 5(2):185-196

Clypeomorus alaseaensis Wissema:
4(1):109

Clypeomorus batillarieformis Habe and
Kosuge: 4(1):109

Clypeomorus bifasciata (Sowerby):
4(1):109

230 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Clypeomorus bifasciata persica ssp. nov.:
4(1):109

Clypeomorus brevis (Quoy and Gaimard,
1834): 4(1):109

Clypeomorus inflata (Quoy and Gaimard,
1834): 4(1):109

Clypeomorus irrorata (Gould): 4(1):109

Clypeomorus nympha nom. nov.: 4(1):109

Clypeomorus pellucida (Hombron and
Jacquinot): 4(1):109

Clypeomorus petrosa (Wood, 1828):
4(1):109

Clypeomorus petrosa chemnitziana
Pilsbry, 1901: 4(1):109

Clypeomorus petrosa gennesi (Fischer
and Vignal): 4(1):109

Clypeomorus purpurastoma Houbrick:
4(1):109

Clypeomorus tjiolonganensis (K. Martin):
4(1):109

Clypeomorus verbeekii (H. Woodward):
4(1):109

Cochlodesma praetenue (Pulteney, 1799):
2:35-40; S1:35-50

Cochlostyla (Hypselostyla) carinata (Lea):
3(1):98-99

Cochlostyla (Orthostylus) pithogaster
(Ferussac): 3(1):98-99

Cochlostyla pithogaster (Ferussac):
3(1):98-99

Codakia orbicularis (Linné, 1758): $1:23-24

Codium: 5(2):259-280

Codium isthmocladium Vickers:
5(2):259-280

Coleoptera: S2:69-81

Collembola: 5(2):185-196

Collisella pelta ‘Rathke’ Escholtz, 1833:
2:80

Collisella scabra Gould, 1846: S1:35-50

Colpidium: S1:79-83

Columbellidae Swainson, 1840: 3(1):96

Concavus Newman, 1982: 4(1):39-42

Concavus finchii (Lea, 1833): 4(1):39-42

Conchifera: 6(1):57-68

Conidae Rafinesque, 1815: 4(1):109-111

Conradilla caelata (Conrad, 1834):
1:43-50; 4(1):25-37; 6(1):19-37

Conus Linné, 1758: 3(1):95; 4(1):109-110;

4(2):229

Conus chrysocestus Berry, 1968:
3(1):63-82

Conus figulinus Linné, 1758: 4(1):185-199
(passim)

Conus jaspideus stearnsi Conrad, 1869:
4(1):185-199 (passim)

Conus marylandicus Green, 1830:
4(1):39-42

Conus poormani Berry, 1968: 3(1):63-82

Conus vicweei Old, 1973: 1:75-78

Convoluta convoluta: S1:35-50

Cophocara Stewart, 1927: 4(2):236

Coralliophila incompta Berry, 1960:
3(1):63-82

Corambe Bergh, 1869: 5(2):243-258

Corambidae Bergh, 1869: 5(2):243-258

Corbicula Mihlfeldt, 1844: 1:96; 2:86;
3(1):85-88, 106-107; 5(1):21-30 (passim);
$2:1-5, 41-45, 47-52, 53-58, 59-61,
63-67, 83-88, 89-94, 95-98, 125-132

Corbicula aegyptica ‘Bourguinat’ Ger-
main, 1907: S2:113-124

Corbicula africana (Krauss, 1848):
$2:113-124

Corbicula agrensis ‘Kurr’ Prime, 1860:
$2:113-124

Corbicula arata ‘Theobald’ Sowerby,
1878: S2:113-124

Corbicula artini Pallary, 1903: S2:113-124

Corbicula astartina Martens, 1860:
$2:113-124

Corbicula aurea Heude, 1880: S2:113-124

Corbicula australis (Lamarck, 1818):
$2:113-124

Corbicula baudoni Morlet, 1886:
$2:113-124

Corbicula bitruncata Martens, 1908:
$2:113-124

Corbicula blandiana Prime, 1864:
$2:113-124

Corbicula bocourti (Morelet, 1865):
$2:113-124

Corbicula colorata Martens, 1905:
$2:113-124

Corbicula cor (Lamarck, 1818):
$2:113-124

Corbicula crocea Temcharoen, 1971:
$2:113-124

Corbicula cunningtoni Smith, 1906:
$2:113-124

Corbicula debilis (Gould,1850): S2:113-124

Corbicula elatior Martens, 1905:
$2:113-124

Corbicula erosa Prime, 1861: S2:113-124

Corbicula felnouilliana Heude, 1880:
$2:113-124

Corbicula ferghanensis Kursalova and
Starobogatov, 1971: S2:113-124

Corbicula fischeri Germain, 1907:
$2:113-124

Corbicula fluminalis (Muller, 1774):
5(1):91-99; S2:113-124, 203-209

Corbicula fluminea (Miller, 1774): 1:13-20,
96, 97, 100; 2:86, 87; 3(1):41-45,
47-53, 94, 100, 100-101, 104-105;
3(2):233-242, 267-268, 269, 272;
4(1):21-23, 61-79, 81-88, 115-116, 116,
116-117; 4(2):234; 5(1):1-7, 31-39,
91-99; 6(2):165-178, 199-206;
$1:35-50, 187-191, 193-201; S2:1-5,
7-39, 69-81, 83-88, 89-94, 99-111,
113-124, 133-142, 143-150, 151-166,
167-178, 179-184, 185, 187-191,
193-201, 203-209, 211-218, 219-222,
223-229, 231-239

Corbicula gubernatoria Prime, 1867:
$2:113-124

Corbicula gustaviana Martens, 1900:
$2:113-124

Corbicula heardi Brandt, 1974: S2:113-124

Corbicula iravadica ‘Blanford’ Hanley and
Theobald, 1876: S2:113-124

Corbicula japonica Prime, 1864: S2:1-5,
113-124

Corbicula javanica (Mousson, 1849):
$2:113-124

Corbicula kirkii Prime, 1864: S2:113-124

Corbicula krishnaea Ray, 1967: S2:113-124

Corbicula lamarckiana Prime, 1864:
$2:113-124

Corbicula largillierti (Philippi, 1844):
$2:113-124

Corbicula larnaudieri Prime, 1862:
$2:113-124

Corbicula leana Prime, 1864: 4(1):81-88;
$2:7-39, 203-209

Corbicula leviuscula Prime, 1864:
$2:113-124

Corbicula ligidana Prime, 1861: S2:113-124

Corbicula lindoensis Bollinger, 1914:
$2:113-124

Corbicula loehensis Kruimel, 1913:
$2:113-124

Corbicula lydigiana Prime, 1861: S2:113-124

Corbicula malaccensis Deshayes, 1854:
$2:113-124

Corbicula manilensis (Philippi, 1844):
1:43-50; 4(1):81-88; S2:1-5, 7-39

Corbicula matanensis Sarasin and
Sarasin, 1898: S2:113-124

Corbicula messageri Bavay and Dautzen-
berg, 1901: S2:113-124

Corbicula moltkiana Prime, 1878: S2:113-124

Corbicula moreletiana Prime, 1867:

$2:113-124

Corbicula nitens (Philippi, 1844):
$2:113-124

Corbicula noetlingi Martens, 1899:
$2:113-124

Corbicula occidentiformis Brandt, 1974:
$2:113-124

Corbicula oliphantensis Craven, 1880:
$2:113-124

Corbicula orientalis (Lamarck, 1818):
$2:113-124

Corbicula papyracea Heude, 1880:
$2:113-124

Corbicula petiti ‘Clessin’ Morlet, 1886:
$2:113-124

Corbicula pingensis Brandt, 1974:
$2:113-124

Corbicula pisidiformis Prime, 1866:
$2:113-124

Corbicula planata Martens?: S2:113-124
Corbicula pulchella (Mousson, 1848):

$2:113-124

Corbicula pullata Philippi, 1850:
$2:113-124

Corbicula purpurea Prime, 1863:
$2:113-124

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 231

Corbicula pusilla (‘Parreys’ Philippi,
1847): S2:113-124

Corbicula radiata (‘Parreys’ Philippi,
1846): S2:113-124

Corbicula regia Clessin, 1879: S2:113-124

Corbicula regularis Prime, 1860:
$2:113-124

Corbicula rivalis (‘Busch’ Philippi, 1850):
$2:113-124

Corbicula sandai Reinhardt, 1878: S2:1-5

Corbicula siamensis Prashad, 1929:
$2:113-124

Corbicula sikorae Ancey, 1890: S2:113-124

Corbicula sinensis nom. dub.: S2:7-39,
113-124

Corbicula solidula Prime, 1860:
$2:113-124

Corbicula squalida Deshayes, 1854:
$2:113-124

Corbicula striatella Deshayes, 1854:
$2:113-124

Corbicula subradiata ‘Kurr’ Prime, 1861:
$2:113-124

Corbicula suifuensis Lindholm, 1925:
$2:113-124

Corbicula sumatrana Clessin, 1887:
$2:113-124

Corbicula tanganyicensis Crosse, 1881:
$2:113-124

Corbicula tenuis Clessin, 1887: S2:113-124

Corbicula tibetensis Prashad, 1929:
$2:113-114

Corbicula tobae Martens, 1900: S2:113-124

Corbicula tumida Deshayes, 1854:

— $2:113-124

Corbicula vinca Heude, 1880: S2:113-124

Corbicula virescens Brandt, 1974:
$2:113-124

Corbicula vokesi Brandt, 1974: S2:113-124

Corbiculacea Gray, 1847: 3(2):201-212;
4(1):116; 5(1):21-30 (passim)

Corbiculidae Gray, 1847: 4(1):116

Cordylophora lacustris Allman:
5(2):287-292

Corona Albers, 1850: 3(1):8 (passim)

Corophium: $3:59-70

Corophium spinicoine: S2:7-39

Corophium stimpsoni: S2:7-39

(Corynadenia) Berry, 1940: 3(1):63-82

Coryphella Gray, 1850: 5(2):185-196

Coryphella gracilis (Alder and Hancock,
1844): 5(2):287-292

Coryphella nobilis Verrill, 1880:
5(2):287-292

Coryphella pellucida (Alder and Hancock,
1843): 5(2):287-292

Coryphella salmonacea (Couthony, 1839):
4(2):205-216; 5(2):287-292 (passim)

Coryphella verrilli Kuzirian: 5(2):287-292

Coryphella verrucosa (Sars, 1829):
5(2):287-292

Cosmetalepas Iredale, 1924: 2:21-34

Costasiella lilanae: 4(2):205-216 (passim)

Costasiella ocellifera (Simroth, 1895):
5(2):197-214, 259-280

Costasiella nonatoi Marcus and Marcus,
1960: 5(2):259-280

Costasiellidae: 5(2):259-280

Cottus carolinae (Gill): 5(1):1-7

Couthouyella Bartsch, 1909: S$1:1-22

Cranchia (Liocranchia) globula Berry,
1909: 3(1):63-82

Cranchioidea Berry, 1920: 3(1):63-82

Crania californica Berry, 1921: 3(1):63-82

Crassatella corbuloides Reeve, 1842: 2:83

Crassatella laevis A. Adams, 1854: 2:83

Crassatella lomiteensis Oldroyd, 1924: 2:83

Crassatella marginata Keep, 1887: 2:83;

3(1):103

Crassatella ponderosa (Gmelin, 1791):
4(2):238

Crassatella vadosa Morton, 1834:
4(2):238

Crassatellidae Férussac, 1822: 4(2):238

Crassatellinae Ferussac, 1822: 2:38

Crassilabrum wittichi (Hertlein and Jor-
dan, 1927): 4(1):1-12

Crassinella nuculiformis Berry, 1940:
3(1):63-82

Crassispira starri Hertlein and Jordan,
1927: 4(1):1-12

Crassostrea Sacco, 1897: 1:35-42,
108-109; 4(2):157-162

Crassostrea angulata (Lamarck, 1819):
4(2):157-162

Crassostrea columbiensis (Hanley, 1846):
4(2):157-162

Crassostrea cortiezensis (Hertlein, 1951):
1:108

Crassostrea gigas (Thurnberg, 1793):
1:102; 4(2):157-162; S2:7-39 (passim)

Crassostrea guyanensis: 1:35-42

Crassostrea lacerta: 1:35-42

Crassostrea rhizophorae (Guilding, 1818):
1:35-42, 102, 108

Crassostrea virginica (Gmelin, 1791):
1:105-106, 108, 109; 2:41-50, 63-73;
3(1):85-88; 4(1):101; 4(2):157-162;
6(2):189-197 (passim); S1:59-78, 79-83,
101-109 (passim), 111-116; S3:1-4, 5-10,
11-16, 17-23, 25-29, 31-36, 37-40, 41-49,
59-70, 71-75

Crassostreinae: 4(2):157-162

Crassostreini: 4(2):157-162

Cratena capensis Barnard, 1927:
5(2):243-258

Cratena peregrina (Gmelin, 1791):
5(2):197-214

Cratena simba Edmunds, 1970:
5(2):243-258

Cratenidae: 5(2):243-258

Crenella Brown, 1827: S1:23-24

Crenimargo Berry, 1963: 3(1):63-82

Crenimargo electis Berry, 1963: 3(1):63-82

Crepidula Lamarck, 1799: 3(1):85-88;
4(1):1-12; 6(1):9-17

Crepidula aculeata (Gmelin, 1791):
4(2):173-183

Crepidula adunca Sowerby, 1825:
3(1):33-40; 4(2):173-183; 6(1):17

Crepidula cerithicola C. B. Adams:
4(2):173-183

Crepidula coei Berry, 1950: 3(1):63-82

Crepidula convexa Say, 1822: 1:110;
3(1):33-40; 4(2):173-183

Crepidula costata Morton, 1829: 4(1):39-42

Crepidula costata Sowerby, 1824:
4(1):39-42

Crepidula dilatata Lamarck, 1822:
4(2):173-183

Crepidula echinus (Broderip, 1834):
4(2):173-183

Crepidula fecunda: 4(2):173-183

Crepidula fornicata (Linné, 1758): 1:110;
3(2):135-142 (passim), 179-186
(passim); 4(2):165-172; 6(1):17;
$1:35-50, 85-90; S2:203-209

Crepidula incurva (Broderip, 1834):
4(2):173-183

Crepidula lessonii (Broderip, 1834):
4(2):173-183

Crepidula lingulata Gould, 1846:
4(2):173-183

Crepidula maculosa Conrad, 1846:
4(2):173-183

Crepidula monoxyla (Lesson, 1830):
4(2):173-183

Crepidula navicula Morch, 1877:
4(2):173-183

Crepidula nummaria Gould, 1846: 3(1):33-40

Crepidula onyx Sowerby, 1824: 1:110;
3(1):33-40; 4(2):173-183, 241-242;
6(1):17

Crepidula philippiana: 4(2):173-183

Crepidula plana Say, 1822: 1:110;
3(1):33-40; 4(2):173-183; S1:85-91

Crepidula protea Orbigny, 1841: 1:110

Crepidula striolata Menke, 1851: 1:110;
4(2):173-183

Crimora Alder and Hancock, 1862:
5(2):243-258

Crimora coneja Marcus, 1961: 5(2):197-214

Crimora papillata Alder and Hancock,
1862: 5(2):185-196, 197-214

Cristaria (Pletholophus) discoidea (Lea):
5(1):91-99 (passim)

Crossaster papposis (Linné, 1758):
5(2):287-292

Croton sp.-09: 1:67-70

Crucibulum castellum Berry, 1963:
3(1):63-82

Crucibulum cyclopium Berry, 1969:
3(1):63-82

Crucibulum inerme Nelson, 1870: 4(1):1-12

Crucibulum marense: 4(2):173-183

Crucibulum monticulus Berry, 1969:
3(1):63-82

Crucibulum personatum Keen, 1958:
4(2):173-183

232 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Crucibulum scutellatum (Wood, 1828):
4(1):1-12; 4(2):173-183

Crucibulum spinosum (Sowerby, 1824):
4(2):173-183, 241-242

Crucibulum subactum Berry, 1963:
3(1):63-82

Crucibulum umbrella (Deshayes, 1830):
4(2):173-183

Cryptoconchus Burrow, 1815: 6(1):79-114

Cryptoconchus floridanus (Dall, 1889):
6(1):79-114

Cryptomphalis (Helix) aspersa (Miller,
1774): 5(2):303-306

Cryptostrea: 4(2):157-162

Cryptostrea permollis (Sowerby, 1871):
4(2):157-162

Cryptostreini: 4(2):157-162

Cryptozona belangeri (Deshayes):
4(1):114; 4(2):237

Ctenodonta nasuta (Hall): 4(1):111-112

Cumanotus beaumonti (Eliot, 1906):
5(2):197-214

Cumberlandia Ortmann, 1912: 4(1):13-19

Cumberlandia monodonta (Say, 1829):
4(1):13-19, 25-37; 6(1):19-37

Curvemysella Habe, 1959: 1:90-91

Curvemysella paula (Adams, 1856): 1:90-91

Cuspidariidae Dall, 1886: S1:35-50

Cuthona Alder and Hancock, 1855:
5(2):243-258

Cuthona adyarensis Rao, 1952:
5(2):197-214

Cuthona albocrusta MacFarland:
5(2):197-214, 287-292

Cuthona albopunctata (Schmekel):
5(2):197-214

Cuthona amoena (Alder and Hancock,
1845): 5(2):185-196

Cuthona annulata (Baba, 1949):
5(2):243-258

Cuthona caerulea (Montagu, 1804):
5(2):197-214

Cuthona cocoachroma Williams and
Gosliner: 5(2):197-214

Cuthona columbiana (O'Donoghue, 1921):
5(2):197-214

Cuthona concinna (Alder and Hancock,
1843): 5(2):185-196, 287-292

Cuthona divae (Marcus, 1961): 5(2):197-214

Cuthona foliata (Forbes and Goodsir,
1839): 5(2):185-196

Cuthona genovae (O’Donoghue, 1929):
5(2):197-214

Cuthona granosa (Schmekel): 5(2):197-214

Cuthona ilonae (Schmekel): 5(2):197-214

Cuthona kanga (Edmunds, 1970):
5(2):243-258

Cuthona kuiteri Rudman: 5(2):185-196

Cuthona ministriata (SGchmekel):
5(2):197-214

Cuthona nana (Alder and Hancock,
1842): 5(2):185-196, 197-214, 287-292

Cuthona ocellata (Schmekel): 5(2):197-214

Cuthona ornata Baba, 1937: 5(2):243-258

Cuthona poritophages Rudman:
5(2):185-196, 197-214

Cuthona pustulata (Alder and Hancock,
1854): 5(2):197-214

Cuthona speciosa (Macnae, 1954):
5(2):243-258

Cyamiacea: 3(1):103

Cyanogaster Blainville, 1825: 5(2):215-241

Cyanophyceae: S2:167-178

Cyanoplax fackenthallae Berry, 1919:
3(1):63-82

Cyclinella Dall, 1902: 4(1):1-12

Cyclocardia borealis (Conrad, 1831):
$1:59-78

Cyclonaias tuberculata (Rafinesque,
1820): 1:29, 43-50, 51-60; 2:85, 85-86;
3(1):105; 4(1):25-37; 6(1):19-37;
6(2):165-178

Cyclonaias tuberculata granifera (Lea,
1838): 6(1):19-37

Cyclonaias tuberculata tuberculata
(Rafinesque, 1820): 6(1):19-37

Cyclophoridae: 3(2):223-231; S1:1-22

Cyclostremella Bush, 1897: S1:1-22

Cyclostremellidae: S1:1-22

Cyerce antillensis Engel, 1927: 5(2):259-280

Cyerce cristallina (Trinchese, 1881):
5(2):197-214

Cylichna cylindracea (Pennant, 1777):
5(2):185-196

Cylichna tubulosa Gould, 1859:
5(2):243-258

Cylichnidae A. Adams, 1850: 4(2):233

Cylinchna Loven, 1846: $1:1-22

Cylinchnella canaliculata (Say, 1826): 1:91

Cylindrobulla P. Fischer, 1856: S1:1-22

Cylindrobullidae Thiele, 1926: 5(2):243-258;
$1:1-22

Cymatioa Berry, 1964: 3(1):63-82

Cymatium nicobaricum (Roding, 1798):
$1:35-50

Cymatium parthenopeum (Von Salis,
1793): S1:85-91

Cymatium perryi Emerson and Old, 1963:
1:75-78

Cymbulia Péron and Lesueuer, 1810:
$1:1-22

Cymbuliidae Gray, 1840: S1:1-22

Cymia chelonia: 2:84-85

Cymia heimi Hertlein and Jordan, 1927:
4(1):1-12

Cymopolia: 5(2):259-280

Cyphoma gibbosum (Linné, 1758): 2:84

Cyrpraea sp.: 2:84

Cypraea amandusi Hertlein and Jordan,
1927: 4(1):1-12

Cypraea talpa (Linne, 1758): 2:84

Cypraecassis testiculus (Linne, 1758):
$1:35-50

Cyprinus carpio: S2:69-81, 89-94

Cyprogenia irrorata (Lea, 1830): 1:29;
4(1):25-37; 6(1):19-37; 6(2):165-178

Cyprogenia stegaria (Rafinesque, 1820):
1:31-34; 2:85-86; 4(1):25-37;
6(1):19-37; 6(2):165-178

Cyrtonaias tampicoensis berlandieri (Lea,
1857): 2:86

Cyrtoplax sykesi Thiele, 1909: 6(1):115-130

Cyrtoplax (Notoplax) speciosa H. Adams,
1861: 6(1):115-130

Dacrydium Torrell, 1859: 4(1):111-112;
$1:23-24

Daphne: 5(2):183-184

Daphnia: S1:79-83

Delphinula trigonostoma Lamarck, 1822:
2:57-61

Dendostrea Sowerby, 1839: 4(2):157-162

Dendostrea folium (Linné, 1758):
4(2):157-162

Dendostrea frons (Linné, 1758):
4(2):157-162

Dendostrea mexicana (Sowerby, 1871):
4(2):157-162

(Dendrochiton) Berry, 1911: 3(1):63-82;
6(1):141-151

Dendrochiton laurae Berry, 1963: 3(1):63-82

Dendrochiton lirulatus Berry, 1963:
3(1):63-82; 6(1):141-151

Dendrochiton psales Berry, 1963:
3(1):63-82

Dendrochiton semiliralatus Berry, 1927:
3(1):63-82

Dendrodorididae Pruvot-Fol, 1935:
5(2):243-258

Dendrodoris Ehrenberg, 1831:
5(2):185-196, 243-258

Denarodoris albopunctata Cooper, 1863:
4(2):205-216 (passim)

Dendrodoris caesia (Bergh, 1907):
5(2):243-258

Dendrodoris denisoni (Angas, 1864):
5(2):243-258

Dendrodoris krebsii (Morch, 1863):
5(2):197-214

Dendrodoris miniata (Alder and Hancock,
1864): 5(2):197-214

Dendrodoris nigra (Stimpson, 1855):
5(2):197-214, 243-258

Dendronotacea Gray, 1857: 5(2):215-241

Dendronotus diversicolor Robilliard, 1970:
5(2):197-214, 287-292

Dendronotus frondosus (Ascanius, 1774):
4(2):205-216 (passim); 5(2):197-214

Dendronotus iris Cooper, 1863:
5(2):197-214

Dentalium Linné, 1758: S1:35-50

Dermatobranchus Hassett, 1824:
5(2):243-258

Dermatobranchus Striatellus Baba, 1949:
5(2):197-214

Deroceras agreste Linne, 1758): 6(1):16

Deroceras carunae (Pollonera, 1891):
6(1):16

Deroceras laeve (Miller, 1774): 1:23
(passim), 110; 6(1):16

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 233

Deroceras reticulatum (Miiller, 1774):
1:110; 3(2):223-231; 6(1):16

Detracia ‘Gray’ Turton, 1840: S1:1-22

Diadumene leucolena (Verrill): S3:59-70

Diala goniochila: 4(2):235

Diaphana Brown, 1837: $1:1-22

Diaphana californica Dall, 1919:
5(2):197-214

Diaphana minuta (Brown, 1827):
5(2):185-196

Diaphanidae Odhner, 1922: S1:1-22

Diaphorodoris \redale and O’Donoghue,
1923: 2:95

Diaphorodoris papillata Portmann and
Sandmeier, 1960: 5(2):185-196

Diastomidae Cossmann, 1895: 4(2):235

Diaulula sandiegensis (Cooper, 1862):
5(2):185-196

Dicta odhneri Schmekel: 5(2):197-214

Dimya californica Berry, 1936: 3(1):63-82

Dimya coralliotis Berry, 1944: 3(1):63-82

Diodora Gary, 1821: 2:21-34

Diodora cayenensis (Lamarck, 1822):
4(1):107-108

Diodora pusilla Berry, 1959: 3(1):63-82

Diadorini: 2:21-34

Dinophycea: S2:167-178

Diplodonta impolita Berry, 1953:
3(1):63-82

Diptera: S2:69-81

Diptychophilia Berry, 1964: 3(1):63-82

Dirona albolineata Cockrell and Eliot,
1905: 5(2):197-214

Dirona aurantia Hurst, 1966: 5(2):197-214

Discodorididae Bergh, 1891: 5(2):243-258

Discodoris Bergh, 1877: 5(2):185-196

Discodoris cavernae Starmiihiner, 1955:
5(2):185-196

Discodoris erythraeensis Vayssiere, 1912:
5(2):197-214

Discodoris fragilis (Alder and Hancock,
1864): 5(2):243-258

Discodoris heathi MacFarland, 1905:
5(2):197-214

Discodoris indecora Bergh, 1881:
5(2):185-196

Discodoris maculosa Bergh, 1884:
5(2):197-214

Discodoris sandiegensis (Cooper, 1863):
5(2):197-214

Disconaias salinasensis (Simpson, 1908):
2:86

Discotectonica Marwick, 1931:
4(1):108-109

Discus (Gonyodiscus?) brunsoni Berry,
1955: 3(1):63-82

Discus rotundatus (Miller, 1776): 3(1):27
(passim)

Divalinga comis (Olsson, 1964): 4(1):1-12

Docoglossa Troschel, 1866: S1:1-22

Donax fossor (Say, 1822): 3(1):92

Donax trunculus (Linné, 1758): 1:13
(passim)

Dondersiidae: 6(1):57-68

Dondice Marcus, 1958: 5(2):183-184

Dondice paguerensis Brandon and
Cutress: 5(2):185-196

Dolabella auricularia (Solander, 1786):
5(2):243-258

Dolabrifera Gray, 1847: 5(2):185-196

Dolabrifera dolabrifera (Range, 1828):
5(2):243-258

Doridacea: 5(2):215-241, 243-258

Doridella obscura Verrill, 1870:
5(2):185-196, 197-214

Doridella steinbergae (Lance, 1962):
5(2):185-196, 197-214

Dorididae Rafinesque, 1815: 5(2):243-258

Doridomorpha gardineri Eliot, 1906:
5(2):185-196

Doriodoxa benthalis Barnard, 1963:
5(2):243-258

Doriopsilla Bergh, 1880: 5(2):185-196,
243-258

Doriopsilla miniata (Alder and Hancock,
1864): 5(2):243-258

Doriopsilla pharpa Marcus, 1961:
5(2):185-196, 197-214

Doriopsis pecten (Collingwood, 1881):
5(2):243-258

Doris Linné, 1758: 5(2):185-196

Doris ocelligera (Bergh): 5(2):197-214

Doris verrucosa Linne, 1758: 5(2):243-258

Doryteuthis plei (Blainville, 1823):
6(2):213-217

Doto acuta Schmekel and Kress, 1977:
5(2):197-214

Doto amyra Marcus, 1961: 5(2):197-214

Doto coronata (Gmelin, 1791):
5(2):197-214, 243-258

Doto doerga Marcus and Marcus, 1963:
5(2):197-214

Doto kya Marcus, 1961: 5(2):197-214

Doto paulinae Trinchese, 1881:
5(2):197-214

Doto pinnatifida (Montague, 1804):
5(2):243-258

Doto rosea Trinchese, 1881: 5(2):197-214,
243-258

Dotoidae Gray, 1853: 5(2):243-258

Dreissena polymorpha Pailas: 5(1):91-99
(passim); S2:124 (passim), 174
(passim), 219-222

Drillia (Clathrodrillia) Dall, 1918: 4(1):1-12

Dromus dromas (Lea, 1834): 1:43-50;
3(1):41-45; 4(1):25-37, 117; 6(1):19-37;
6(2):165-178

Dromus dromas caperatus (Lea, 1845):
6(1):19-37

Dromus dromas dromas (Lea, 1834):
6(1):19-37

Dugesia tigrina: S2:7-39, 89-94

Durvilledoris leminiscata (Quoy and
Gaimard, 1832): 5(2):243-258

Dysnomia arcaeformis (Lea, 1831):
6(1):19-37

Dysnomia biemarginata (Lea, 1857):
1:43-50

Dysnomia brevidens (Lea, 1834): 1:43-50;
6(1):19-37

Dysnomia capsaeformis (Lea, 1834):
1:43-50; 6(1):19-37

Dysnomia flexuosa: 6(1):19-37

Dysnomia florentina (Lea, 1857): 1:43-50;
6(1):19-37

Dysnomia florentina walkeri (Wilson and
Clark, 1914): 6(1):19-37

Dysnomia haysiana (Lea, 1833): 1:43-50;
6(1):19-37

Dysnomia lenior (Lea, 1842): 6(1):19-37

Dysnomia lewisi (Walker, 1910): 6(1):19-37

Dysnomia stewardsoni (Lea, 1852):
6(1):19-37

Dysnomia sulcata delicata: 3(1):105

Dysnomia torulosa (Rafinesque, 1820):
1:43-50; 6(1):19-37

Dysnomia torulosa gubernaculum (Reeve,
1865): 6(1):19-37

Dysnomia torulosa propinqua (Lea, 1857):
6(1):19-37

Dysnomia torulosa rangiana (Lea, 1839):
3(1):105

Dysnomia triquetra (Rafinesque, 1820):
1:43-50; 3(1):105; 6(1):19-37

Dysnomia turgida (Lea, 1848): 6(1):19-37

Ebala: S1:1-22

Elaeocyma baileyi Berry, 1969: 3(1):63-82

Elaeocyma ricaudae Berry, 1969: 3(1):63-82

Electra crustulenta (Pallas): 5(2):185-196;
197-214

Electra pilosa (Linné): 4(1):103-104;
5(2):197-214, 293-301

Eledone cirrhosa (Lamarck): 4(2):217-227;
6(1):45-48

Eledone moschata: 4(2):217-227

Eledonella heathi Berry, 1911: 3(1):63-82

Eledonella pygmaea Verrill, 1848:
4(2):217-227

Elimia sp.: 4(1):25-37

Elimia potosiensis (Lea): 3(1):100

Elimina sp.: 1:31-34

Ellipsaria lineolata (Rafinesque, 1820):
2:85-86; 4(1):25-37; 6(1):19-37

Elliptio Rafinesque, 1820: 1:109-110;
5(2):125-128; 6(2):165-178

Elliptio angustata (Lea, 1831): 1:95

Elliptio angustatus Lea, 1831: 3(1):94

Elliptio (Canthyria) steinstansana Johnson
and Clarke: 3(1):104-105

Elliptio cistelliformis (Lea, 1863): 1:61-68

Elliptio complanata (Lightfoot, 1786):
1:109-110; 3(1):104-105; 5(1):31-39

Elliptio crassidens (Lamarck, 1819): 1:29,
43-50, 109-110; 3(1):41-45, 47-53;
4(1):21-23, 25-37; 6(1):19-37;
6(2):165-178

Elliptio crassidens crassidens (Lamarck,
1819): 1:51-60; 2:85-86; 4(1):117;
5(2):165-171

234 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Elliptio dilatata (Rafinesque, 1820): 1:29,
31-34, 51-60; 2:85-86; 3(1):41-45,
47-53, 105; 4(1):27-37, 117;
5(2):165-171; 6(1):19-37; 6(2):165-178

Elliptio dilatatus (Rafinesque, 1820):
1:43-50; 6(1):19-37

Elliptio dilatatus delicatus (Simpson):
5(2):165-171

Elliptic emmonsii Lea, 1857: 3(1):94

Elliptio fisheriana (Lea, 1838): 1:61-68

Elliptio fisherianus Lea, 1838: 3(1):94

Elliptio foliculatus Lea, 1838: 3(1):94

Elliptio folliculata (Lea, 1838): 1:61-68

Elliptio hazelhurstianus Lea, 1858: 3(1):94

Elliptio icterina (Conrad, 1834): 1:95;
4(1):117; 4(2):231

Elliptio lanceolata (Lea, 1820): 1:61-68,
94-95, 95, 109-110; 3(1):94;
$2:203-209

Elliptio producta (Conrad, 1836): 1:61-68

Elliptio productus Conrad, 1836: 3(1):94

Elliptio ravenelli (Conrad, 1834): 1:61-68

Elliptio shepardiana (Lea, 1834): 3(1):94

Elliptio subcylindraceus Lea, 1873: 3(1):94

Elliptio waccamawensis (Lea, 1863):
1:61-68

Elliptoideus Frierson, 1927: 1:109-110

Ellobiidae H. and A. Adams, 1855: S1:1-22

Ellobium Roding, 1798: S1:1-22

Elodea: 6(2):179-188 (passim)

Elysia Risso, 1818: 5(2):287-292; $1:1-22

Elysia arena Carlson and Hoff:
5(2):185-196

Elysia cauze Marcus, 1957: 4(2):205-216
(passim)

Elysia chlorotica (Gould, 1870):
5(2):197-214

Elysia flava Verrill, 1901: 5(2):259-280

Elysia furvicauda Burn: 5(2):259-280

Elysia halimedae Macnae, 1954:
5(2):243-258

Elysia hedgpethi (Marcus, 1961):
5(2):197-214

Elysia hopei (Marcus): 5(2):197-214

Elysia livida Baba, 1955: 5(2):243-258

Elysia marginata (Pease, 1871):
5(2):243-258

Elysia moebii (Bergh, 1888): 5(2):243-258

Elysia olivaceus: 4(1):109-111

Elysia papillosa Verrill, 1901: 4(2):232;
5(2):259-280

Elysia patina Marcus: 5(2):197-214

Elysia rufescens (Pease, 1871):
5(2):243-258

Elysia subornata Verrill, 1901: 4(2):232;
5(2):197-214, 259-280

Elysia timida (Risso, 1818): 5(2):197-214

Elysia tuca Marcus, 1967: 4(2):232;
5(2):197-214, 259-280

Elysia vatae Risbec, 1928: 5(2):243-258

Elysia virgata (Bergh, 1888): 5(2):243-258

Elysia viridis (Montagu, 1804):
5(2):243-258

Elysiidae: 4(2):232; 5(2):243-258,
259-280; S1:1-22

Emarginulinae Gray, 1834: 2:21-34

Embletonia Alder and Hancock, 1851:
5(2):185-196

Embletonia fuscata Gould, 1870:
4(2):205-216 (passim)

Embletonia gracilis Risbec, 1928:
5(2):243-258

Embletonia pulchra Alder and Hancock,
1844: 5(2):303-306

Embletonia pulchra faurei (Alder and Han-

cock): 5(2):197-214

Embletoniidae: 5(2):243-258

Endodontidae: 2:97; 5(2):243-258

Enigmonia aenigmatica (Holton):
5(2):159-164 (passim)

Enoploteuthis galaxias Berry, 1918:
3(1):63-82

Enoploteuthoidea Berry, 1920: 3(1):63-82

Enoptroteuthis Berry, 1920: 3(1):63-82

Enoptroteuthis spinicauda Berry, 1920:
3(1):63-82

Ensis Schumacher, 1817: 2:96

Ensis directus Conrad, 1843: S1:59-78

Ensis myrae Berry, 1953: 3(1):63-82

Entodesma Philippi, 1845: S1:35-50

Entomotaeniata: S1:1-22

Eolidina mannarensis Rao and
Alagarswami, 1960: 5(2):197-214

Ephadra Gistel, 1848: 2:21-34

Ephemeroptera: S2:69-81

Epimenia verrucosa (Nierstrasz):
6(1):57-68

Epioblasma Rafinesque, 1831:
4(1):117-118; 6(2):165-178

Epioblasma arcaeformis (Lea, 1831):
4(1):25-37; 6(1):19-37; 6(2):165-178

Epioblasma biemarginata (Lea, 1857):
6(1):19-37

Epioblasma brevidens (Lea, 1834):
4(1):25-37; 6(1):19-37; 6(2):165-178

Epioblasma capsaeformis (Lea, 1831):
4(1):25-37; 6(1):19-37; 6(2):165-178

Epioblasma flexuosa (Rafinesque, 1820):
4(1):25-37, 117; 6(1):19-37

Epioblasma florentina (Lea, 1857):
4(1):25-37; 6(2):165-178

Epioblasma florentina florentina (Lea,
1857): 6(1):19-37; 6(2):165-178

Epioblasma florentina walkeri (Wilson and
Clark, 1914): 6(1):19-37

Epioblasma haysiana (Lea, 1834):
3(1):41-45; 4(1):25-37; 6(1):19-37;
6(2):165-178

Epioblasma lenior (Lea, 1842): 6(1):19-37

Epioblasma lewisi (Walker, 1910):
4(1):25-37; 6(1):19-37

Epioblasma obliquata (Rafinesque, 1820):
4(1):25-37

Epioblasma propinqua (Lea, 1857):
4(1):25-37; 6(1):19-37

Epioblasma rangiana (Lea, 1839): 1:31-34

——

Epioblasma sampsoni (Lea, 1861):
1:27-30, 31-34

Epioblasma stewardsoni (Lea, 1852):
4(1):25-37; 6(1):19-37; 6(2):165-178

Epioblasma sulcata (Lea, 1824):
4(1):25-37; 6(1):19-37

Epioblasma torulosa (Rafinesque, 1820):
6(1):19-37

Epioblasma torulosa cincinnatiensis (Lea,
1840): 6(1):19-37

Epioblasma torulosa gubernaculum
(Reeve, 1865): 4(1):25-37; 6(1):19-37

Epioblasma torulosa torulosa (Rafinesque,
1820): 2:85-86; 4(1):25-37; 6(1):165-178

Epioblasma triquetra (Rafinesque, 1820):
1:29; 4(1):25-37; 6(1):19-37

Epioblasma turgidula (Lea, 1848):
6(1):19-37

Epiphragmorpha petricola Berry, 1916:
3(1):63-82

Epiphragmorpha petricola orotes Berry,
1920: 3(1):63-82

Epiphragmorpha petricola sangabrielis
Berry, 1920: 3(1):63-82

Epiphragmorpha traskii chrysoderma
Berry, 1920: 3(1):63-82

Epiphragmorpha traskii willetti Berry,
1920: 3(1):63-82

Epiphragmorpha tudiculata allyniana
Berry, 1920: 3(1):63-82

Epiphragmorpha tudiculata rufiterrae
Berry, 1916: 3(1):63-82

Epitoniacea Berry, 1910: S1:1-22

Epitoniidae: Berry, 1910: $1:1-22

Epitonium Roding, 1798: $1:1-22

Epitonium albidum (Orbigny, 1842): 1:1-12;
3(1):47-53, 85-88; 4(1):185-199
(passim)

Epitonium greenlandicum (Perry, 1811): 1:1
(passim)

Epitonium millecostatum (Pease,
1860-1861): 1:1, 2, 9 (passim)

Epitonium rupicola (Kurtz, 1860): 1:1, 7, 9
(passim)

Epitonium tinctum (Carpenter, 1864): 1:1,
5 (passim)

Epitonium ulu Pilsbry, 1921

Ercolania funerea (Costa): 5(2):197-214,
259-280

Ercolania fuscata (Gould): 5(2):197-214,
259-280

Escherichia coli: 3(2):179-186

Eubranchidae Odhner, 1934: 5(2):243-258

Eubranchus Forbes, 1838: 5(2):243-258

Eubranchus coniclus: 5(2):183-184

Eubranchus exiguus (Alder and Han-
cock): 5(2):185-196, 197-214

Eubranchus farrani (Alder and Hancock,
1848): 5(2):185-196, 197-214

Eubranchus olivaceus (O’Donoghue,
1922): 5(2):197-214

Eubranchus rustyus (Marcus, 1961):
5(2):197-214

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 235

Eubranchus sanjuanensis Roller:
5(2):287-292

Eubranchus tricolor (Forbes, 1838):
5(2):287-292

Euchelus gemmatus (Gould, 1895):
4(2):232-233

Eucleoteuthis Berry, 1916: 3(1):63-82

Eucrassinella Cruz, 1980: 2:83

Eucrassinella fluctuata (Carpenter, 1864):
2:83

Eucrassatella Iredale, 1924: 2:83

Eucrassatella aequitorialis Cruz, 1980: 2:83

Eucrassatella antillarum (Reeve, 1842): 2:83

Eucrassatella digueti (Lamy, 1917): 2:83

Eucrassatella gibbosa (Sowerby, 1832): 2:83

Eucrassatella (Hybolophus) gibbosa tucilla
Olsson, 1932: 2:83

Eucrassatella manabiensis Cruz, 1980: 2:83

Eudoxochiton nobilis Gray, 1843:
6(1):141-151

Euglandia rosea: 2:98-99

Euglenophycaea: S2:167-178

Euhadra: 2:97

Eukiefferiella sp.: 3(2):151-168

Eulimacea: $1:1-22

Eulimidae: S1:1-22

Eunicella verrucosa (Pallas): 5(2):185-196

Eupera cubensis: $2:223-229

Euplera caudata (Say, 1822): 4(1):185-199
(passim); S3:59-70

Eupleura caudata etterea Baker, 1951:
2:63-73

Euplica turturina (Lamarck, 1822):
4(2):232-233

Euprymna: 4(2):217-227

Euprymna scolopes Berry, 1913:
3(1):63-82

Eurycaelon anthonyi: 4(1):25-37

Eurypanopeus depressus (Smith):
$3:59-70

Eurystomella bilabriata Hincks:
5(2):185-196

Euselenops Pilsbry, 1896: 5(2):215-241

Euselenops luniceps (Cuvier, 1817):
5(2):215-241, 243-258

Eutheostoma flabellare Rafinesque: 5(1):1-7

Eutheostoma rufilineatum (Cope): 5(1):1-7

Euthyneura Spengel, 1881: S1:1-22

Facelina bostoniensis (Couthony, 1838):
5(2):287-292

Facelina coronata (Forbes and Goodsir,
1839): 5(2):185-196

Facelina dubia Pruvot-Fol, 1948:
5(2):197-214

Facelina fusca Schmekel: 5(2):197-214

Facelina olivacea Macnae, 1954:
5(2):243-258

Facelina punctata (Alder and Hancock,
1864): 5(2):197-214

Facilinidae Bergh, 1889: 5(2):243-258

Falcidens: 6(1):57-68; S1:23-34

Fargoa bartschi (Winkley, 1909): S1:1-22

Fasciolaria tulipa (Linné, 1758): 4(1):113

Fasciolariidae Gray, 1853: 4(1):109-110

Favartia garretti (Pease, 1869): 2:84

Favorinus branchialis (Rathke):
5(2):185-196

Favorinus ghanensis Edmunds, 1968:
5(2):243-258

Favorinus japonicus Baba, 1949:
5(2):243-258

Ferrissia Walker, 1903: 5(1):73-84

Ferrissia fragilis (Tryon, 1863): 3(1):99;
5(1):9-19

Ferrissia parallela (Haldeman, 1844):
5(1):9-19

Ferrissia rivularis (Say, 1819): 3(2):135-142,
243-265; 5(1):105-124 (passim)

Ferrissia wautieri: 3(2):151-168

Fimbria fimbriata (Linné, 1758): 5(1):21-30
(passim)

Fiona pinnata (Eschscholtz, 1831):
5(2):197-214, 243-258

Fionidae Gray, 1857: 5(2):243-258

Fissurelidea annulus Odhner, 1932: 2:21-34

Fissurella aperta Sowerby, 1825: 2:21-34

Fissurella hiantula Lamarck, 1822: 2:21-34

Fissurella minosti Melleville, 1843: 2:21-34

Fissurellidaea (Sic) bimaculata Dall, 1871:
2:21-34

Fissurellidea Orbigny, 1841: 2:21-34

Fissurellidea bimaculata Dall, 1871: 2:21-34

Fissurellidea hiantula Pilsbry, 1890: 2:21-34

Fissurellidea megatrema Orbigny, 1841:

2:21-34

Fissurellidea patagonica (Strebel, 1907):
2:21-34

Fissurellidea patagonicus (Strebel, 1907):
2:21-34

Fissurellidea (Pupillaea) aperta (Sowerby,
1825): 2:21-34

Fissurellidini Pilsbry, 1890: 2:21-34

Fissurellinae Fleming, 1822: 2:21-34

Flabella fuscus (O’Donoghue, 1924):
5(2):197-214

Flabella salmonacea (Couthouy, 1838):
5(2):197-214

Flabella trilineata (O'Donoghue, 1921):
5(2):197-214

Flabella verrucosa (Sars, 1829):
5(2):197-214

Flabellina Voight, 1834: 5(2):243-258

Flabellina affinis (Gmelin, 1791):
5(2):197-214

Flabellina capensis (Thiele, 1925):
5(2):243-258

Flabellina funeka Gosliner and Griffiths,
1981: 5(2):243-258

Flabellinidae Bergh, 1889: 5(2):243-258

Fontelicella: 4(2):243

Fossaria modicella (Say, 1825): 3(1):99

Fragilaria: S2:167-178

Fucus serratus (Linné, 1758):
5(2):293-301

Fucus vesiculosus 1:92

Fundulus: 2:1-20

Fusconaia Simpson, 1900: 1:109-110;
6(2):165-178

Fusconaia barnesiana (Lea, 1838):
1:43-50; 3(1):41-45, 104; 4(1):25-37;
6(1):19-37; 6(2):165-178

Fusconaia barnesiana barnesiana (Lea,
1838): 6(1):19-37

Fusconaia barnesiana bigbyensis (Lea,
1841): 1:43-50; 3(1):41-45; 5(1):1-7;
6(1):19-37; 6(2):165-178

Fusconaia barnesiana tumescens (Lea,
1845): 6(1):19-37; 6(2):165-178

Fusconaia cor (Conrad, 1834): 6(2):179-188

Fusconaia cor analoga (Ortmann, 1918):
6(1):19-37

Fusconaia cor cor (Conrad, 1834):
6(1):19-37

Fusconaia cuneolus (Lea, 1840): 1:43-50;
6(1):19-37; 6(2):179-188

Fusconaia cuneolus appressa (Lea, 1871):
6(1):19-37

Fusconaia cuneolus cuneolus (Lea, 1840):
6(1):19-37

Fusconaia ebena (Lea, 1831): 1:51-60;
4(1):117-118; 5(2):177-179; 6(1):19-37,
49-54

Fusconaia edgariana (Lea, 1840):
1:43-50; 3(1):104, 106; 6(1):19-37

Fusconaia edgariana analoga (Ortmann,
1918): 6(1):19-37

Fusconaia flava (Rafinesque, 1820): 1:28,
29, 31-34, 51-60; 3(1):47-53, 93, 105;
4(1):21-23; 5(2):165-171; 6(1):19-37

Fusconaia lateralis (Rafinesque, 1820):
6(1):19-37

Fusconaia maculata maculata (Rafinesque,
1820): 1:31-34; 2:85-86

Fusconaia ozarkensis (Call, 1887): 2:85

Fusconaia pilaris (Lea, 1840): 6(2):165-178

Fusconaia polita Say, 1834: 6(1):19-37

Fusconaia polita lesueriana (Lea, 1840):
6(1):19-37

Fusconaia polita pilaris (Lea, 1840):
6(1):19-37

Fusconaia pusilla (Rafinesque, 1820):
6(1):19-37

Fusconaia subrotunda (Lea, 1831):
1:43-50; 3(1):41-45, 105; 4(1):25-37,
117; 6(1):19-37; 6(2):165-178

Fusconaia subrotunda leseuriana (Lea,
1840): 6(1):19-37

Fusconaia subrotunda pilaris (Lea, 1840):
6(1):19-37

Fusconaia subrotunda subrotunda (Lea,
1831): 6(1):19-37

Fusconaia undata (Barnes, 1823):
6(1):19-37

Fuscosaria lineolata (Rafinesque, 1820):
1:51-60

Fusinus acanthodes (Watson, 1882):
3(1):101-102

Fusinus (Pagodula) acanthodes (Watson,
1882): 3(1):101-102

236 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Fusinus pumilus Lea, 1833: 4(1):39-42

Fusiturricula Woodring, 1928: 1:92

Fusus acanthodes (Watson, 1882):
3(1):101-102

Fusus kingii Gabb, 1864: 4(2):236

Gafrarium pectinatum (Linné, 1758):
5(1):91-99 (passim)

Galeomma (Lepirodes?) mexicanum
Berry, 1959: 3(1):63-82

Galiteuthis phyllura Berry, 1911: 3(1):63-82

Gambusia affinis: S2:69-81

Garamella: 5(2):243-258

Gasterosteus aculeatus (Linné, 1758):
5(2):185-196

Gastroheayle: 5(2):281-286

Gastroplax Blainville, 1819: 5(2):215-241

Gastropoda, Unspecified: 1:99, 99-100;
2:80-81, 87-88; 3(1):93, 93-94, 95;
4(1):102-103, 103, 114; 4(2):243, 244;
5(1):101-104; S2:69-81

Gastropteridae Swainson, 1840: 4(2):233;
5(2):243-258

Gastropteron alboaurantium Gosliner,
1984: 5(2):243-258

Gastropteron flavobrunneum Gosliner,
1984: 5(2):243-258

Gastropteron rubrum (Rafinesque, 1814):
5(2):185-196

Gegania Jeffreys, 1884: S1:1-22

Geitodoris capensis Bergh, 1907:
5(2):243-258

Gelonia erosa Berry, 1911: 3(1):33-40

Gemma gemma (Totten, 1834): 2:96

Gemmula hindsiana Berry, 1958: 3(1):63-82

Geukensia demissa (Dillwyn, 1817):
3(1):33-40; 4(2):233-234; S1:59-78

Geukensia demissa demissa (Dillwyn,
1817): 5(1):173-176

Geukensia demissa granosissima (Sower-
by, 19147): 3(1):103; 4(1):112;
5(1):173-176

Gibbula marmorea (Pease, 1867):
4(2):232-233

Gigantonotum Guangyu and Si, 1965:
5(2):215-241

Glaucidae Menke, 1828: 5(2):243-258

Glaucus atlanticus (Forster, 1777):
5(2):185-196, 243-258

Gleba: $1:1-22

Glossiphona complanata: 3(2)151-168;
5(1):73-84

Glossodoris atromarginata (Cuvier, 1804):
5(2):243-258

Glossodoris bilineata Pruvot-Fol, 1953:
5(2):197-214

Glossodoris gracilis von Rapp, 1827:
5(2):197-214

Glossodoris luteopunctata Gantes, 1962:
5(2):197-214

Glossodoris sp.: 5(2):243-258

Glycera: 2:96

Glyptosoma pilsbryanum Berry, 1938:
3(1):63-82

Glyptosoma pilsbryanum binneyanum
Berry, 1938: 3(1):63-82

Godiva quadricolor (Barnard, 1929):
5(2):243-258

Gonatopsis borealis Sasaki, 1923:
2:89-90

Gonatus berryi Naef, 1923: 2:89

Gonatus madokai (Berry, 1921): 2:89

Gonatus magister Berry, 1913: 3(1):63-82

Gonatus middendorfi: 4(2):241

Gonatus onyx Young, 1972: 2:89

Gonatus tinro: 2:89

Gonaxis kibweziensis: 2:98-99

Gonaxis quadrilateralis: 2:98-99

Goniobasis sp.: 1:31-34; S2:203-209

Goniobasis albanyensis Lea, 1864: 6(1):17

Goniobasis atherni Clench and Turner:
6(1):17

Goniobasis curvicostata (Reeve, 1861):
6(1):17

Goniobasis dickensoni Clench and
Turner: 6(1):17

Goniobasis floridensis (Reeve, 1860):
6(1):17

Goniobasis laqueata: 1:43-50

Goniobasis proxima (Say, 1825): 1:105;
3(1):99-100

Goniobasis vanhyningiana Goodrich:
6(1):17

Goniodoridae H. and A. Adams, 1854:
5(2):243-258

Goniodoris castanea (Alder and Hancock,
1854): 5(2):197-214, 243-258

Goniodoris mercurialis Macnae, 1958:
5(2):243-258

Goniodoris ovata Barnard, 1934:
5(2):243-258

Gourmya gourmyi (Crosse, 1861): 2:1-20

Granosolarium Sacco, 1892: 4(1):108-109

Granulina oviformis (Orbigny, 1841):
4(1):185-199

Graptacme calamus Dall, 1899: 1:100

Graptemys pulchra Baur: S2:7-39

Gryphaeidae: 4(2):157-162

Gulo: 5(2):183-184

Gymnodinium veneficum: S2:167-178

Gymnodorididae Odhner, 1941:
5(2):243-258

Gymnodoris alba (Bergh, 1877):
5(2):243-258

Gymnodoris bicolor (Alder and Hancock,
1864): 5(2):243-258

Gymnodoris ceylonica (Kelaart, 1858):
5(2):243-258

Gymnodoris inornata (Bergh, 1880):
5(2):243-258

Gymnodoris limaciformis (Eliot, 1908):
4(1):109-110

Gymnodoris okinawae Baba, 1936:
5(2):243-258

Gymnodoris striata (Eliot, 1904):
5(2):197-214

Gymnosomata Blainville, 1824: $1:1-22

Gymnotoplax Pilsbry, 1896: 5(2):215-241

Gymnotoplax americanus (Verrill, 1885):
5(2):215-241

Gyraulus Charpentier, 1837: 2:88

Gyraulus circumstriatus (Tryon, 1866):
3(1):99; 5(1):9-19

Gyraulus deflectus (Say, 1824): 3(1):99;
5(1):9-19

Gyraulus parvus (Say, 1817): 5(1):9-19,
31-39, 73-84

Haematopus bachmani: 2:80

Haemopsis grandis (Verrill): 5(1):73-84

Halgerda formosa Bergh, 1880:
5(2):243-258

Halgerda punctata Farran, 1905:
5(2):243-258

Halgerda wasinensis Eliot, 1904:
5(2):243-258

Halichondria panicea (Pallas):
5(2):185-196, 197-214

Halimeda discoidea: 5(2):259-280

Halimeda incrassata: 5(2):259-280

Halimeda simulans: 5(2):259-280

Haliotis cracherodii Leach, 1814:
4(2):233-234

Haliotis corrugata Wood, 1828:
3(2):223-231

Haliotis roei (Gray, 1827): 3(1):97

Haliotis rufescens Swainson, 1822:
3(2):223-231

Halisarca dujardini Johnston: 4(1):103-104

Hallaxa apefae: 5(2):183-184

Hallaxa chani Gosliner and Williams:
5(2):197-214

Halodakra salmonea (Carpenter, 1832):
2:83; 3(1):103

Halodakra subtrigona (Carpenter, 1857):
3(1):103

Halodule wrighti Ashers, 1868:
4(2):185-199

Halomenia gravida Heath: 6(1):57-68

Haminoea Turton and Kingston, 1830:
5(2):185-196; S1:1-22

Haminoea alfredensis Bartsch, 1915:
5(2):243-258

Haminoea antillarum (Orbigny, 1841):
5(2):197-214

Haminoea hydatis (Linné, 1758):
5(2):185-196

Haminoea natalensis (Krauss, 1848):
5(2):243-258

Haminoea navicula (Da Costa):
5(2):185-196

Haminoea solitaria (Say, 1822):
5(2):197-214

Haminoea vesicula Gould, 1855:
4(2):165-172; 5(2):197-214

Haminoeidae Pilsbry, 1895: 5(2):243-258

Hancockia californica MacFarland, 1923:
5(2):287-292

Hancockia ucinata Hesse,1872:
5(2):197-214

Hanetia capitanea Berry, 1957: 3(1):63-82

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Hanetia macrospira Berry, 1957: 3(1):63-82

Hanetia mendozana Berry, 1959: 3(1):63-82

Hanleya spicata Berry, 1919: 3(1):63-82

Hapalochlaena maculosa (Hoyle):
6(2):207-211

Haplochromis burtoni Giinther:
5(2):185-196

Haplosporidia nelsoni (Haskins, Stauber
and Mackin): S3:5-10

Haplosporidium: S1:101-109; S3:5-10

Haplosporidium costalis Wood and
Andrews: S3:59-70

Haplosporidium (Minchinia) nelsoni
(Haskins, Stauber and Mackin):
$3:17-23, 59-70

Haplotrematidae Baker, 1931: 1:97

Haustellum wilsoni D’Attilio and Old, 1971:
1:75-78

Hedylopsidae: $1:1-22

Hedylopsis Thiele, 1931: 5(2):281-286;
$1:1-22

Hedylopsis spiculifera (Kowalevsky, 1901):
5(2):303-306

Heliaucus Orbigny, 1842: $1:1-22

Heliaucus cylindricus (Gmelin, 1871):
$1:1-22

Heliacus (Grandeliacus) \redale, 1957:
4(1):108-109

Heliacus (Gyriscus) Tiberi, 1867:
4(1):108-109

Heliacus (Heliacus) Orbigny, 1842:
4(1):108-109

Heliaucus perreieri (Rochebrune, 1881):
$1:1-22

Heliacus (Teretropoma) Rochebrune, 1881:
4(1):108-109

Heliacus (Torinista) |redale, 1936:
4(1):108-109

Helicinidae Gray, 1842: 3(2):223-231

Heliococranchia fisheri Berry, 1901:
3(1):63-82

Helicostylinae: 3(1):98-99

Heliopora: 5(2):185-196

Helisoma Swainson, 1840: S1:51-58

Helisoma anceps (Menke): 3(1):99;
4(1):118-119; 5(1):9-19, 31-39, 73-84,
105-124 (passim)

Helisoma campanulatum (Say, 1821):
3(1):99; 5(1):9-19

Helisoma duryi Weatherby: S1:35-50

Helisoma trivolvis (Say, 1817):
3(2):213-221, 243-265; 4(1):118-119;
4(2):229; 5(1):9-19; 6(1):17

Helix Linné, 1758: 4(2):157-162 (passim);
$1:35-50

Helix aspersa Miller, 1774: 1:24, 97-98;
3(1):27 (passim); 6(1):16; S1:35-50

Helix pomacea Linne, 1758: 1:97-98;
6(1):16; S1:35-50

Helix pomatia Linné, 1758: 3(2):223-231

Helix vulgaris: 1:13 (passim)

Helminthoglypta arrosa humboldtica
Berry, 1935: 3(1):63-82

Helminthoglypta ayersiana (Newcomb,
1861): 3(1):103

Helminthoglypta (Charodotes) traskii
(Newcomb, 1861): 3(1):103

Helminthoglypta crotalina Berry, 1928:
3(1):63-82

Helminthoglypta dupetithouarsii consors
Berry, 1938: 3(1):63-82

Helminthoglypta euomphalodes Berry,
1938: 3(1):63-82

Helminthoglypta graniticola Berry, 1926:

3(1):63-82

Helminthoglypta inglesi Berry, 1938:
3(1):63-82

Helminthoglypta isabella Berry, 1938:
3(1):63-82

Helminthoglypta jaegeri Berry, 1928:
3(1):63-82

Helminthoglypta liodoma Berry, 1938:
3(1):63-82

Helminthoglypta mohaveana Berry, 1926:
3(1):63-82

Helminthoglypta napea Berry, 1938:
3(1):63-82

Helminthoglypta orina Berry, 1938:
3(1):63-82

Helminthoglypta proles saccharodytes
Berry, 1938: 3(1):63-82

Helminthoglypta riparia Berry, 1928:
3(1):63-82

Helminthoglypta tejonis Berry, 1938:
3(1):63-82

Helminthoglypta thermimontis Berry, 1953:

3(1):63-82

Helminthoglypta traskii (Newcomb), 1861):
3(1):103

Helminthoglypta tudiculata angelena
Berry, 1938: 3(1):63-82

Helminthoglypta tudiculata kernensis
Berry, 1930: 3(1):63-82

Helminthoglypta tularensis pluripuncta
Berry, 1938: 3(1):63-82

Helminthoglyptidae Pilsbry, 1939: 1:97;
2:98; 3(1):8 (passim), 103

Hemistena lata (Rafinesque, 1820):
6(1):19-37; 6(2):165-178

Hemitrochus: 3(1):8 (passim)

Hendersonia occulta (Say, 1831): 1:99

Hermaea bifida (Montagu, 1816):
5(2):197-214

Hermissenda crassicornis (Eschscholtz,
1831): 1:13 (passim); 4(2):205-216;
5(2):287-292

(Herpeteros) Berry, 1947: 3(1):63-82

Heterobranchia: S1:1-22

Heterodonta Neumayr, 1884: 4(1):111-112

Heterogastropoda: $1:1-22

Heteroglossa: S1:1-22

Heteroterma Gabb, 1869: 4(2):236

Hertleinella Berry, 1958: 3(1):63-82

Hertleinella leucostephes Berry, 1958:
3(1):63-82

Hexabranchidae: 5(2):243-258

237

Hexabranchus marginatus: 5(2):185-196

Hexabranchus sanguineus (Riippell and
Leuckart, 1828): 5(2):185-196,
243-258

Hexaplex erythrostomus (Swainson, 1831):
6(1):45-48

Hiatella Bosc, 1801: 3(2):135-142 (passim)

Hindsia nodulosa (Whiteaves, 1874):
4(2):236

Hipponix grayanus Menke, 1853:
4(2):173-183

Hipponix pilosus (Deshayes, 1832):
4(1):1-12

Hopkinsia rosacea MacFarland, 1905:
5(2):185-196

Hoplodoris nodulosa (Angas, 1864):
5(2):197-214

Hormospira Berry, 1958: 3(1):63-82

Hyalina avena (Kiener, 1834): 4(1):185-199
(passim)

Hybolophus Stewart, 1930: 2:83

Hydatina Schumacher, 1817: 5(2):185-196;
$1:1-22

Hydatina albocincta (van der Hoeven,
1811): 5(2):243-258

Hydatina amplustre (Linne, 1758):
5(2):243-258

Hydatina physis (Linne, 1758):
5(2):243-258

Hydatina zonata (Lightfoot, 1786):
5(2):243-258

Hydatinidae Pilsbry, 1895: 5(2):243-258;
$1:1-22

Hyaractinia echinata Fleming:
5(2):185-196, 287-292

Hyadrobia truncata: 4(1):101-102

Hydrobia ulvae (Pennant): S1:35-50

Hydrobiidae Stimpson, 1865:
3(2):223-231; 4(2):243

Hydrocenidae: 3(2):223-231

Hydroides dianthus (Verrill, 1873): S1:1-22
(passim)

Hydropsyche: S2:69-81

Hyotissa Stenzel, 1971: 1:90;
4(2):157-162

Hyotissa hyotis (Linné, 1758): 4(2):157-162

Hyotissini: 4(2):157-162

Hypselodoris bennetti (Angas, 1864):
5(2):197-214

Hypselodoris bilineata (Pruvot-Fol, 1953):
5(2):185-196

Hypselodoris cantabrica Bouchet and
Ortega: 5(2):185-196

Hypselodoris capensis (Barnard, 1927):
5(2):243-258

Hypselodoris carnea (Bergh, 1889):
5(2):243-258

Hypselodoris gracilis (Rapp, 1827):
5(2):185-196

Hypselodoris infucata (RUppel and
Leuckart, 1828): 5(2):243-258

Hypselodoris maridadilus Rudman, 1977:
5(2):243-258

238 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Hypselodoris messinensis (von Ihering,
1880): 5(2):185-196, 197-214

Hypselodoris tema Edmunds: 5(2):185-196

Hypselodoris valenciennesi (Cantraine,
1841): 5(2):185-196

Hypselodoris webbi (Orbigny, 1839):
5(2):185-196

Hypselodoris zebra (Heilprin, 1888):
5(2):185-196

(Hypselostyla): 3(1):98-99

Ictalurus furcatus (Lesueur): S2:7-39,
89-94

Ictalurus punctatus: S2:69-81, 89-94,
211-218

Ictiobus bubalus (Rafinesque): S2:7-39,
89-94

Ictiobus cyprinellus (Valenciennes):
$2:7-39

Ictiobus niger (Rafinesque): S2:7-39, 89-94

Idasola |Iredale, 1915: S1:23-34

Idasola argentea (Jeffreys, 1876): S1:23-34

Idiosepius: 4(2):217-227

Idiosepius notoides Berry, 1921: 3(1):63-82

Illex Steenstrup, 1880: 4(2):217-227

Illex coindetii (Verany, 1837): S1:93-100

Illex illecebrosus (Lesueur, 1821): 1:90;
2:51-56; 3(1):107; 4(1):55-60, 101;
4(2):239, 240-241; S1:93-100

Illex oxygonius Roper, Lu and Mangold,
1969: S1:93-100

llyanassa obsoleta (Say, 1822): 2:14
(passim); 4(1):110; 4(2):165-172;
6(2):189-197 (passim); S1:35-50

lo fluvialis (Say, 1825): 4(1):25-37;
5(1):65-72 (passim); 6(2):165-178

lo verrucosa lima: 1:43-50

Ischnochiton Gray, 1847: 6(1):115-130

Ischnochiton herdmani: 6(1):141-151

Ischnochiton haersoltei Kaas, 1954:
6(1):115-130

Ischnochiton kilburni Kaas, 1979:
6(1):115-130

Ischnochiton luzonicus (Sowerby, 1842):
6(1):115-130

Ischnochiton ranjhai Kaas, 1954:
6(1):115-130

Ischnochiton rissoi (Payraudeau): 6(1):57-68

Ischnochiton rufopunctatus Odhner, 1919:
6(1):115-130

Ischnochiton sansibarensis Thiele, 1910:
6(1):115-130

Ischnochiton striolatus (Gray, 1828):
4(1):107-108

Ischnochiton winckworthi Leloup, 1936:
6(1):115-130

Ischnochiton yerbury Smith, 1891:
6(1):115-130

Ischnochiton (lschnochiton) winckworthi
Leloup, 1936: 6(1):115-130

Ischnochiton (Ischnichiton) yerburyi
(Smith, 1891): 6(1):115-130

Ischnochiton (Lepidozona) amabilis Berry,
1917: 3(1):63-82

Ischnochiton (Lepidozona) asthenes
Berry, 1919: 3(1):63-82

Ischnochiton (Lepidozona) californiensis
Berry, 1931: 3(1):63-82

Ischnochiton (Lepidozona) gallina Berry,
1925: 3(1):63-82

Ischnochiton (Lepidozona) golischi Berry,
1919: 3(1):63-82

Ischnochiton (Lepidozona) interfossa
Berry, 1917: 3(1):63-82

Ischnochiton (Lepidozona) luzonicus
(Sowerby, 1842): 6(1):115-130

Ischnochiton (Lepidozona) nipponica
Berry, 1918: 3(1):63-82

Ischnochiton (Lepidozona) pilsbryanus
Berry, 1917: 3(1):63-82

Ischnochiton (Lepidozona) sanc-
taemonicae Berry, 1922: 3(1):63-82

Ischnochiton (Lepidozona) willetti Berry,
1917: 3(1):63-82

Ischnochiton (Radsiella) delagoaensis
Ashby, 1931: 6(1):115-130

Ischnochitonidae Dall, 1889: 6(1):115-130

Ischnochitoninae Pilsbry, 1893: 6(1):115-130

Isochrysis: S1:85-91

Isochrysis galbana (Parke): 6(2):189-197

Isochrysis galbiana (Parke): 3(1):33-40;
4(1):81-88, 89-99

lsonychia: S2:69-81

Janolidae: 5(2):243-258

Janolus capensis Bergh, 1907:
5(2):243-258

Janolus longidentatus Gosliner, 1981:
5(2):243-258

Janthina sp.: 1:4, 7, 9, 10; $1:1-22

Janthina exigua Lamarck, 1816: S1:1-22

Janthina janthina (Linné, 1758): S1:1-22,
35-50

Janthinidae Leach, 1823: S1:1-22

Joannisia Monterosato, 1884: 5(2):215-241

Joculator ridicula Watson, 1866:
4(2):232-233

Jorunna tormentosa (Cuvier, 1804):
4(1):103-104; 5(2):185-196, 243-258

Jorunna zania Marcus, 1976: 5(2):243-258

Joubiniteuthis Berry, 1920: 3(1):63-82

Julia exquisita Gould, 1862: 4(2):232-233

Julia zebra Kawaguti, 1981: 5(2):243-258

Juliamitrella: 3(1):96

Juliidae Smith, 1885: 5(2):243-258;
$1:1-22

Kalinga ornata Alder and Hancock, 1864:
5(2):243-258

Kaloplocamus ramosus (Contraine, 1835):
5(2):243-258

Katharina tunicata Wood, 1815: 6(1):141-151

Kellia rosea Dall, Bartsch and Rehder,
1938: 4(2):232-233

Kermia aniani Kay, 1979: 4(2):232-233

Kentrodorididae: 5(2):243-258

Kirchenpaueria pinnata (Linne):
5(2):197-214

Knefastia Dall, 1919: 4(1):1-12

Knefastia princeps Berry, 1953: 3(1):63-82

Knefastia walkeri Berry, 1953: 3(1):63-28

Koloonella hawaiiensis Kay, 1979:
4(2):232-233

Koonsia Verrill, 1882: 5(2):215-241

Lacuna cossmanni: $1:23-24

Lacuna succinea Berry, 1953: 3(1):63-82

Lacuna vincta (Montagu, 1803):
5(2):287-292

Laetmoteuthis Berry, 1916: 3(1):63-82

Laetmoteuthis lugubris Berry, 1913:
3(1):63-82

Laevapex fuscus (C. B. Adams): 3(1):99;
3(2):243-265 (passim); 5(1):9-19,
105-124 (passim)

Laevicardium substriatum (Conrad, 1837):
4(2):241-242

Laevicaulis alte (Férussac): $1:35-50

Laicus argentatus Pontopidan:
5(2):185-196

Lalia cockerelli MacFarland, 1905:
5(2):197-214, 287-292

Lamellaria perspicua (Linné, 1758):
4(1):185-199 (passim)

Lamellibranchia: S1:23-34

Lamellidens Simpson, 1900: 4(1):13-19

Laminaria saccharina (Linné): 5(2):185-196

Lampadioteuthidae Berry, 1916: 3(1):63-82

Lampadioteuthis Berry, 1916: 3(1):63-82

Lampadioteuthis megaleia Berry, 1916:
3(1):63-82

Lamprotula leai (Gray): 5(1):91-99

Lampsilis Rafinesque, 1820: 1:109-110;
4(1):13-19, 117-118; 6(2):165-178;
$1:35-50

Lampsilis abrupta Say, 1831: 6(1):19-37

Lampsilis altilis (Conrad, 1834): 1:94

Lampsilis anodontoides (Lea, 1834): 1:29,
43-50; 6(1):19-37

Lampsilis anodontoides fallaciosa (Smith,
1899): 6(1):19-37

Lampsilis anodontoides floridensis (Lea,
1852): S2:7-39

Lampsilis cardium cardium (Rafinesque,
1820): 6(1):19-37

Lampsilis cardium satura (Lea, 1852):
6(1):19-37

Lampsilis claibornensis (Lea, 1838):
3(2):233-242; 4(1):21-23; $2:7-39

Lampsilis crocata (Lea, 1841): 1:61-68

Lampsilis fasciola Rafinesque, 1820:
1:43-50; 2:85-86; 3(1):41-45, 47-53,
104, 105; 4(1):25-37; 5(1):1-7;
6(1):19-37; 6(2):165-178

Lampsilis higginsi (Lea, 1857): 1:51-60;
4(2):230; 6(1):39-43, 49-54

Lampsilis ochracea (Say, 1816): 1:61-68;
3(1):104-105

Lampsilis orbiculata (Hildreth, 1836): 2:85,
85-86; 4(1):25-37; 6(1):19-37

Lampsilis ovata (Say, 1817): 1:29, 43-50;
2:85-86; 3(1):41-45, 93, 104;
4(1):25-37; 6(1):19-37; 6(2):165-178

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 239

Lampsilis ovata satura (Lea, 1852):
6(1):19-37

Lampsilis ovata ventricosa (Barnes, 1823):
1:43-50; 4(1):21-23; 6(1):19-37;
6(2):165-178

Lampsilis perovalis (Conrad, 1834): 1:94

Lampsilis radiata (Gmelin, 1792): 3(1):93;
4(1):13-19; 5(1):31-39

Lampsilis radiata luteola (Lamarck, 1819):
1:51-60; 2:85-86, 86; 3(1):47-53, 105;
4(1):21-23; 4(2):230-231; 5(2):165-171

Lampsilis radiata siliquoidea (Barnes,
1823): 1:29; 6(1):39-43

Lampsilis reeviana (Lea, 1852): 2:85

Lampsilis siliquoida (Barnes, 1823):
6(1):19-37

Lampsilis straminea claibornensis (Lea,
1838): 4(1):21-23

Lampsilis teres (Rafinesque, 1820): 2:86;
6(1):19-37

Lampsilis teres anodontoides (Lea, 1831):
1:51-60; 4(1):21-23; 5(2):165-171

Lampsilis teres teres (Rafinesque, 1820):
1:51-60, 71-74; 4(1):117; 5(2):165-171;
6(1):19-37

Lampsilis uniominatus (Simpson, 1900):
$2:7-39

Lampsilis ventricosa (Barnes, 1823): 1:18
(passim), 31-34, 51-60; 2:85-86;
3(1):47-53, 105; 5(2):165-171;
6(1):39-43

Lampsilis virescens (Lea, 1858): 6(1):19-37

Laomedea: 5(2):185-196, 197-214

Laomedea loveni: 5(2):197-214

Lapsigyrus Berry, 1958: 3(1):63-82

Lasaeidae Gray, 1847: 1:90-91

Lasmigona Rafinesque, 1831: 6(2):165-178

Lasmigona badia (Rafinesque, 1831):
6(1):19-37

Lasmigona complanata (Barnes, 1823):
1:43-50, 51-60, 71-74; 3(1):105;
4(1):117-118; 5(2):165-171; 6(1):19-37

Lasmigona compressa (Lea, 1829):
3(1):98, 105; 5(2):165-171

Lasmigona costata (Rafinesque, 1820):
1:29, 43-50, 51-60; 2:35-40, 82,
85-86; 3(1):47-53, 104, 105;
4(1):25-37, 117-118; 5(2):165-171,
6(1):19-37; 6(2):165-178

Lasmigona holstonia (Lea, 1838):
6(1):19-37; 6(2):165-178

Lasmigona subviridis (Conrad, 1835):
2:85-86; 6(2):179-188

Lasmigona undulatus undulatus (Say,
1817): 4(1):117-118

Lastena lata (Rafinesque, 1820): 1:43-50;
6(1):19-37

Laternula Roding, 1798: 2:35-40

Laternula truncata (Lamarck, 1818): 3(1):104

Laternulidae Hedley, 1918: 2:35-40

Latia: $1:1-22

Latiidae: $1:1-22

Laurencia johnstonii: 5(2):185-196

Laurencia obtusa Lamouroux, 1813:
4(2):185-199

Laurencia poitei Lamouroux, 1813:
4(2):185-199

Lecithophorus capensis Macnae, 1958:
5(2):243-258

Leiosolenus Carpenter, 1856: 1:101

Leiostomus xanthurus (Lacépede:
$3:59-70

Leminda millecra Griffiths, 1985:
5(2):243-258

Lemindidae: 5(2):243-258

Lemiox rimosa (Rafinesque, 1820):
4(1):25-37; 6(1):19-37

Lemiox rimosus Rafinesque, 1831:
6(1):19-37; 6(2):165-178

Lepetidae Dall, 1869: 4(1):115

Lepidochiton cinereus (Linné, 1767):
6(1):131-139

Lepidochitona cinerea (Linné, 1767):
6(1):57-68, 69-78, 153-159

Lepidochitona corrugata Reeve: 6(1):57-68

Lepidochitona dentiens (Gould, 1846):
6(1):141-151

Lepidochitona flectens (Carpenter, 1864):
6(1):141-151

Lepidochitona keepiana Berry, 1948:
3(1):63-82

Lepidochitona dentiens (Gould, 1846):
4(2):243

Lepidochitonidae Dall, 1899: 6(1):141-151

Lepidopleurus asellus (Gmelin, 1791):
6(1):69-78

Lepidopleurus bottae Rochebrune, 1882:
6(1):115-130

Lepidopleurus cajetanus Poli, 1791:
6(1):131-139, 141-151, 153-159

Lepidopleurus cancellatus (Sowerby,
1839): 6(1):69-78

Lepidopleurus rochebruni Jousseaume,
1893: 6(1):115-130

Lepidopleurus (Xiphiozona) heathi Berry,
1919: 3(1):63-82

Lepidozona Pilsbry, 1892: 6(1):115-130

Lepidozona inefficax Berry, 1963:
3(1):63-82

Lepidozona luzonica (Sowerby, 1842):
6(1):115-130

Lepidozona pella Berry, 1963: 3(1):63-82

Lepidozona retiporosa: 6(1):141-151

Lepidozona subtilis Berry, 1956: 3(1):63-82

Lepidozona (Lepidozona) luzonica (Sower-
by, 1842): 6(1):115-130

Lepomis gibbosus (Linné): 5(1):73-84

Lepomis macrochirus: S2:69-81

Lepomis microchirus: S2:89-94

Lepomis microlophus (Gunther):
5(1):73-84; S2:7-39, 89-94

Leptaxinus Verrill and Bush, 1898: 2:96

Leptaxinus minutus Verrill and Bush,
1898: 2:96

Leptochiton clarkicax Berry, 1922:
3(1):63-82

Leptochiton diomedeae Berry, 1917:
3(1):63-82

Leptodea Rafinesque, 1820: 4(1):117-118

Leptodea fragilis (Rafinesque, 1820): 1:29,
43-50, 51-60, 71-74; 2:85-86; 3(1):105;
4(1):21-23, 25-37; 5(2):165-171;
6(1):19-37

Leptodea leptodon (Rafinesque, 1820):
1:71-74; 3(1):105; 6(1):19-37

Leptonacea Gray, 1847: 1:90-91

Leptosynapta: 2:96

Leptothyra rubricincta (Highels, 1845):
4(2):232-233

Leptothyra verruca (Gould, 1845):
4(2):232-233

Leptoxis arkansensis (Hinkley): 3(1):100

Leptoxis (Athearnia) crassa (Haldeman,
1841): 4(1):25-37

Leptoxis carinata (Bruguiere):
3(2):169-177, 269-272; 4(1):119

Leptoxis crassa anthonyi (Redfield, 1854):
4(1):25-37

Leptoxis praerosa (Say, 1824): 4(1):25-37;
6(2):165-178

Leptoxis subglossa (Say, 1825):
4(1):25-37; 6(2):165-178 (passim)

Leucophyta bidentata: 3(1):27-32

Leucophytia: S1:1-22

Leuroplas McLean, 1970: 2:21-34

Lexingtonia dolabelloides (Lea, 1840):
1:43-50; 3(1):41-45, 104; 4(1):25-37,
104; 6(1):19-37; 6(2):165-178

Lexingtonia dolabelloides conradi (Lea,
1834): 1:43-50

Lexingtonia dolabelloides conradi (Vanat-
ta, 1915): 6(1):19-37

Lienardia baltreata (Pease, 1860):
4(2):232-233

Ligumia nasuta (Say, 1817): 3(1):104-105

Ligumia recta (Lamarck, 1819): 1:29,
51-60; 2:85-86; 3(1):105; 4(1):25-37, 117;
5(2):165-171; 6(1):39-43; 6(2):165-178

Ligumia recta latissima (Rafinesque,
1831): 6(1):19-37

Ligumia subrostrata (Say, 1831):
3(2):233-242; 5(1):41-48; 6(1):19-37;
$1:51-58

Liguus fasciatus (Muller, 1774): 1:98;
3(1):1-10; 5(2):153-157

Liguus fasciatus alternatus Simpson,
1920: 3(1):1-10

Liguus fasciatus aurantius Clench, 1929:
3(1):1-10; 5(2):153-157

Liguus fasciatus barbouri Clench, 1929:
3(1):1-10; 5(2):153-157

Liguus fasciatus beardi Jones, 1979:
3(1):1-10

Liguus fasciatus capensis Simpson, 1920:
3(1):1-10

Liguus fasciatus castanezonatus Pilsbry,
1912: 3(1):1-10; 5(2):153-157

Liguus fasciatus castaneus Simpson,
1920: 3(1):1-10

240 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Liguus fasciatus cingulatus Simpson,
1920: 3(1):1-10

Liguus fasciatus clenchi Frampton, 1932:
3(1):1-10; 5(2):153-157

Liguus fasciatus crassus Simpson, 1920:
3(1):1-10

Liguus fasciatus crenatus ‘Swainson’
Pilsbry, 1912: 3(1):1-10

Liguus fasciatus deckerti Clench, 1935:

3(1):1-10

Liguus fasciatus delicatus Simpson, 1920:
3(1):1-10

Liguus fasciatus dohertyi Pflueger, 1934:
3(1):1-10

Liguus fasciatus dryas Pilsbry, 1932: 3(1):1-10

Liguus fasciatus eburneus Simpson,
1920: 3(1):1-10

Liguus fasciatus elegans Simpson, 1920:
3(1):1-10; 5(2):153-157

Liguus fasciatus elliottensis Pilsbry, 1912:
3(1):1-10

Liguus fasciatus evergladenensis Jones,
1979: 3(1):1-10

Liguus fasciatus farnumi Clench, 1929:
3(1):1-10

Liguus fasciatus floridanus Clench, 1929:
3(1):1-10; 5(2):153-157

Liguus fasciatus framptoni Jones, 1979:
3(1):1-10

Liguus fasciatus fuscoflamellus Frampton,
1932: 3(1):1-10

Liguus fasciatus gloriasylvaticus Doe,
1937: 3(1):1-10

Liguus fasciatus graphicus Pilsbry, 1912:
3(1):1-10

Liguus fasciatus humesi Jones, 1979:
3(1):1-10

Liguus fasciatus innomillatus Pilsbry,
1930: 3(1):1-10

Liguus fasciatus kennethi Jones, 1979:
3(1):1-10

Liguus fasciatus lignumvitae Pilsbry, 1912:
3(1):1-10

Liguus fasciatus lineolatus Simpson,
1920: 3(1):1-10

Liguus fasciatus livingstoni Simpson,
1920: 3(1):1-10; 5(2):153-157

Liguus fasciatus lossmanicus Pilsbry,
1912: 3(1):1-10; 5(2):153-157

Liguus fasciatus lucidovarius Doe, 1937:
3(1):1-10; 5(2):153-157

Liguus fasciatus luteus Simpson, 1920:
3(1):1-10

Liguus fasciatus margaretae Jones, 1979:
3(1):1-10

Liguus fasciatus marmoratus Pilsbry,
1912: 3(1):1-10

Liguus fasciatus matecumbensis Pilsbry,
1912: 3(1):1-10

Liguus fasciatus miamiensis Simpson,
1920: 3(1):1-10; 5(2):153-157

Liguus fasciatus mosieri Simpson, 1920:
3(1):1-10; 5(2):153-157

Liguus fasciatus nebulosus Doe, 1937:
3(1):1-10

Liguus fasciatus ornatus Simpson, 1920:
3(1):1-10; 5(2):153-157

Liguus fasciatus osmenti Clench, 1929:
3(1):1-10

Liguus fasciatus pictus (Reeve, 1842):
3(1):1-10

Liguus fasciatus pseudopictus Simpson,
1920: 3(1):1-10

Liguus fasciatus roseatus Pilsbry, 1912:
3(1):1-10; 5(2):153-157

Liguus fasciatus septentrionalis Pilsbry,
1912: 3(1):1-10

Liguus fasciatus simpsoni Pilsbry, 1921:
3(1):1-10

Liguus fasciatus solida Say, 1825:
3(1):1-10

Liguus fasciatus solidulus Pilsbry, 1912:
3(1):1-10

Liguus fasciatus solidus (Say, 1825):
3(1):1-10

Liguus fasciatus solisocassus DeBoe,
1933: 3(1):1-10

Liguus fasciatus splendidus Frampton,
1932: 3(1):1-10

Liguus fasciatus subcrenatus Pilsbry,
1912: 3(1):1-10

Liguus fasciatus testudineus Pilsbry, 1912:

3(1):1-10; 5(2):153-157

Liguus fasciatus vacaensis Simpson,
1920: 3(1):1-10

Liguus fasciatus versicolor Simpson,
1920: 3(1):1-10

Liguus fasciatus violaftumosus Doe, 1937:
3(1):1-10

Liguus fasciatus vonpaulseni Young,
1960: 3(1):1-10

Liguus fasciatus walkeri Clench, 1933:
3(1):1-10; 5(2):153-157

Liguus fasciatus wintei Humes, 1954:
3(1):1-10

Limacia clavigera (Muller, 1776):
5(2):243-258

Limacinidae Blainville, 1823: $1:1-22

Limapontia Johnston, 1836: S1:1-22

Limapontia capitata (Miiller): 5(2):197-214,
259-280

Limapontiidae Gary, 1847: $1:1-22

Limax marginatus Miller, 1774: 6(1):16

Limax maxima Linné, 1758: 2:78

Limax maximus Linné, 1758: 6(1):16

Limax pseudoflavus Evans: 3(2):223-231;
6(1):16

Limenandra nodosa Haefelfinger and
Stamm: 5(2):197-214

Limifossor: 6(1):57-68

Limnodrilus: S2:7-39

Limnoperna Rochebrune, 1882:
5(2):159-164 (passim)

Limnoperna fortuei (Dunker): 5(1):91-99

Limnoperna lacustris Martens: 5(1):91-99
(passim)

Limnoperna supoti Brandt, 1974:
5(1):91-99 (passim)

Limulus polyphemus: 2:96; 3(1):33-40;
6(1):69-78 (passim)

Liolophura gaimardi: S1:79-83

Lirularia Dall, 1909: 4(1):109

Lirularia lirulata (Carpenter, 1864): 4(1):109

Lissarca notocadensis Mellvill and
Standen: 4(2):235

Lithasia geniculata (Haldeman, 1840):
4(1):25-37

Lithasia geniculata salebrosa (Conrad,
1834): 4(1):25-37

Lithasia obovata (Say, 1829): 1:31-34

Lithasia pinguis (Lea, 1852): 1:27-30

Lithasia verrucosa (Rafinesque, 1820):
4(1):25-37

Lithasia verrucosa lima (Simpson, 1900):
1:43-50

Lithasia (Angitrema) verrucosa (Raf-
inesque, 1820): 6(2):165-178

Lithophaga Roding, 1798: 1:101

Lithophaga lithophaga (Linné, 1758):
6(1):131-139

Lithophaga (Labis) attenuata rogersi
Berry, 1957: 3(1):63-82

Lithophaga nigra (Orbigny, 1842): 1:101

Litiopa Rang, 1829: 4(2):235

Litiopidae: 4(2):235

Littorina Ferussac, 1822: 1:108-109;
6(1):9-17; S1:1-22; S2:203-209

Littorina arcana Ellis: 6(1):17

Littorina filosa: 4(1):112

Littorina irrorata (Say, 1822): 2:78;
3(2):223-231; S1:35-50

Littorina littorea (Linné, 1758): 1:92;
3(1):33-40; 3(2):135-142 (passim);
5(1):105-124

Littorina mespillium (Muhlfeld, 1824):
4(1):185-199

Littorina obtusata (Linné, 1758): 1:92;
4(1):108

Littorina rudis (Dautzenberg and Fisher):
6(1):17

Littorina saxatilis (Olivi, 1792): 1:92-93;
3(1):1-10

Littorina scabra (Linné, 1758): 4(1):112

Littorina ziczac (Gmelin, 1791): 4(2):233

Littorinidae Gray, 1840: 4(2):157-162 (passim)

Lobiger serradifalci (Calcara, 1840):
5(2):197-214

Lobiger souverbiei Fischer, 1856:
2:185-196; 5(2):185-196, 243-258,
259-280;

Lobiger viridis Pease, 1863: 5(2):185-196

Loliginoidea Berry, 1920: 3(1):63-82

Loligo Schneider, 1784: S1:93-100

Loligo brasiliensis Blainville, 1823:
6(2):213-217

Loligo etheridgei Berry, 1918: 3(1):63-82

Loligo forbesi: 4(2):240

Loligo opalescens Berry, 1911: 2:93;
3(1):63-82; 4(1):55-60, 241; 4(2):240

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 241

Loligo peali Lesueur, 1821: 4(1):101;
6(2):213-217

Loligo sanpaulensis Brakoniecki, 1984:
6(2):213-217

Loligo vulgaris Lamarck, 1799: 4(1):55-60;
4(2):217-227

Loliolopsis Berry, 1929: 3(1):63-82

Loliolopsis chiroctes Berry, 1929:
3(1):63-82

Lolliguncula brevis (Blainville, 1823): 1:90;
2:91; 6(2):213-217

Lopha Roding, 1798: 4(2):157-162

Lopha cristagalli (Linné, 1758: 4(2):157-162

Lophinae Vyalov, 1936: 4(2):157-162

Lophini: 4(2):157-162

(Lophochiton) Berry, 1925: 3(1):63-82

Lophocochlia minutissimus (Pilsbry, 1921):
4(2):232-233

Lophopleurella capensis (Thiele, 1912):
5(2):243-258

Lorica (Solivaga) finschi (Thiele, 1910):
6(1):115-130

(Lotoria) Emerson and Old, 1963: 1:75-78

Lottia gigantea (Sowerby, 1834): 2:80;
4(2):242-243; S1:35-50

Lucapinella Pilsbry, 1890: 2:21-34

Lucapinella milleri Berry, 1959: 3(1):63-82

(Lucilina) Dall, 1882: 6(1):115-130

Lucina atlantis McLean, 1936: S1:23-34

Lucina (Linga) pennsylvanica (Linne,
1758): $1:23-34

Lucina (Lucinisca) Dall, 1901: 4(1):1-12

Lucina (Phacoides) pectinatus (Gmelin,
1791): $1:23-34

Lucinidae Fleming, 1828: S1:23-34

Lucinoma Dall, 1901: S1:23-34

Lucinoma atlantis (McLean, 1936):
$1:23-34

Lucinoma filosa Stimpson, 1851: S1:23-34

Lunaia Berry, 1964: 3(1):63-82

Lunaia lunaris Berry, 1964: 3(1):63-82

Lunatia heros (Say, 1822): 3(1):33-40

Lunatia lewisii (Gould, 1847): S1:35-50

Lycoteuthidae Berry, 1914: 3(1):63-82

Lymacina: $1:1-22

Lymnaea (Stagnicola) elodes (Say, 1821):
1:67-70; 3(2):143-150, 213-221,
269-272; 5(1):73-84, 105-124;
6(1):9-17

Lymnaea emarginata (Say, 1821): 5(1):73-84

Lymnaea palustris (Binney, 1865):
3(2):213-221; S1:35-50

Lymnaea peregra (Miller, 1774): 3(1):27-32
(passim); 3(2):135-142 (passim);
5(1):65-72, 73-84, 105-124 (passim)

Lymnaea stagnalis (Linné, 1758): 1:13;
2:78; 3(2):135-142 (passim), 223-231;
5(1):65-72, 73-84; S1:35-50, 51-58

Lyogyrus granum (Say): 5(1):9-19

Lyonsia Turton, 1822: S1:35-50

Lyonsia californica Conrad, 1837:
5(1):173-176 (passim)

Lyonsia floridana (Conrad, 1849): 2:41-50

Lyonsia hyalina Conrad, 1831: 3(1):104

Lyonsiidae Fischer, 1887: S1:35-50

Lyria guildingii (Sowerby, 1844):
3(1):101-102

Lysinoe: 3(1):102-103

Lysinoe ghiesbreghti: 3(1):102-103

Lythophyta: 2:82

Macfarlandaea Marcus and Gosliner,
1984, Syn. Nov.: 5(2):215-241

Macoma Leach, 1819: 1:108-109

Macoma balthica (Linne, 1758): 1:90;
3(2):213-221; 5(1):21-30 (passim);
$1:59-78; S2:7-39

Macoma calcarea (Gmelin, 1791): 2:94

Macrochisma Sowerby, 1839: 2:21-34

Macron hartmanni Hertlein and Jordan,
1927: 4(1):1-12

Mactra clathrodon Lea, 1833: 4(1):39-42

Mactra modicella (Conrad, 1833):
4(1):39-42

Mactra subcuneata Conrad, 1838:
4(1):39-42

Mactra (Mactra) williamsi Berry, 1960:
3(1):63-82

Magilidae: 3(1):11-26

Magnonaias nervosa (Rafinesque, 1820):

1:31-34, 51-60; 4(1):117-118;
4(2):230-231

Malletiidae: 4(1):111-112

Malleus Lamarck, 1799: 4(2):157-162
(passim)

Mancinella Link, 1807: 4(1):110

Mancinella alouina: 4(1):110

Maraunibina verrucosa (Challis):
5(2):281-286

Margarites (Lirularia) aresta Berry, 1941:
3(1):63-82

Margaritifera hembeli (Conrad, 1838):
3(2):233-242

Margaritifera laevis (Haas, 1910):
5(2):125-128

margaritifera margaritifera (Linné, 1758):
4(1):13-19; 5(1):91-99 (passim);
105-124 (passim); 5(2):125-128;
6(2):179-188 (passim)

Margaritifera marrianae: 4(1):13-19

Margaritiferidae Haas, 1940: 4(1):13-19

Marginella aureocincta Stearns, 1872:
4(1):185-199

Marianina rosea Pruvot-Fol, 1930:
5(2):243-258

Marianinidae: 5(2):243-258

Marinula: $1:1-22

Marioniopsis cyanobranchiata (Ruppell
and Leuckart, 1831): 5(2):243-258

Marisa cornuarietis: 3(2):223-231

Mathilda: $1:1-22

Mathildidae: $1:1-22

Maxacteon: $1:1-22

Mazatlania aciculata: 1:92

Medionidus conradicus (Lea, 1834):
1:43-50; 3(1):41-45, 104; 5(1):1-7;
6(1):19-37; 6(2):165-178, 179-188

Megalocranchia pardus Berry, 1916:
3(1):63-82

Megalodonta beckii: 5(1):73-84

Megalonaias Utterback, 1915: 1:109-110

Megalonaias gigantea (Barnes, 1923):
1:29, 43-50; 2:86; 6(1):19-37

Megalonaias nervosa (Rafinesque, 1820):
2:85-86; 4(1):117; 5(2):165-171;
6(1):19-37

Megapallifera mutabilis: 4(2):238

Megatebennus Pilsbry, 1890: 2:21-34

Megatebennus bimaculatus Pilsbry, 1890:
2:21-34

Megatebennus paragonicus Strebel, 1907:
2:21-34

Megathura Pilsbry, 1890: 2:21-34

Megatebhura crenulata (Sowerby, 1825):
2:21-34

Meghimatium: 4(2):238

Meiomenia: 5(2):281-286

Meiopriapulus fijiensis Morse, 1981:
5(2):281-286

Melampidae Stimpson, 1851: S1:1-22

Melampus Montfort, 1810: $1:1-22

Melampus bidentatus Say, 1822:
3(1):27-32 (passim); 3(2):135-142
(passim); 4(1):110-111, 121-122;
4(2):236-237

Melampus califonianus Berry, 1964:
3(1):63-82

Melampus mousleyi Berry, 1964:
3(1):63-82

Melania lineolata Griffith and Pidgeon,
1934: 2:20

Melaniidae: 3(2):223-231

Melanitta fusca (Linné): S3:59-70

Melanitta nigra (Linné): S3:59-70

Melanoclamys: 5(2):243-258

Melanochlamys diomedea (Bergh, 1894):
5(2):197-214

Melanoides tuberculata (Miller):
5(1):105-124; 6(1):17

Melanoposidae: 3(2):223-231

Melanopsis: 2:1-20; 5(1):85-90

Melarpha cincta (Quoy and Gaimard,
1833): 4(1):185-199 (passim)

Melibe fimbriata Alder and Hancock,
1864: 5(2):197-214

Melibe leonina (Gould, 1852): 5(2):197-214

Melibe litvedi Gosliner, 1987: 5(2):243-258

Melibe pilosa Pease, 1860: 5(2):243-258

Melibe rosea Rang, 1829: 5(2):243-258

Mellanella sp.: 2:83

Melongena melongena Linné, 1758:
4(1):1-12

Melongena melongena consors (Sowerby,
1850): 2:84-85; 4(1):1-12

Melongenidae Gill, 1867: 4(2):233

Melosira: S2:167-178

Membranipora: 5(2):185-196

Membranipora crustulenta Pallas:
5(2):185-196

Membranipora villosa Hincks: 5(2):197-214

242 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Mercenaria Schumacher, 1817: 2:96;
3(1):85-88

Mercenaria mercenaria (Linné, 1758):
1:107; 4(1):111; 4(2):149-155; S1:35-50,
59-78; S3:41-49

Mercuria confusa (Frauenfeld): 5(1):85-90

Mercurua punica Letourneux and
Bourguignat): 5(1):85-90

Mesochaetopterus alipes Monroe, 1933:
1:91

Mesodon ‘Rafinesque’ Férussac, 1821:
2:97-98

Mesodon clausus (Say, 1821): 1:97-98

Mesoaon elevatus (Say, 1821): 1:97-98; 2:98

Mesodon (megasoma sp.?) eritrichius
Berry, 1939: 3(1):63-82

Mesodon (megasoma sp.?) euthales
Berry, 1939: 3(1):63-82

Mesodon thyroidus (Say, 1816): 1:97-98

Mesodon zaletus (Binney, 1837): 1:98;
2:97-98, 98

Mesogastropoda Thiele, 1929:
3(2):223-231; S1:1-22, 23-34

Metachaetoderma: 6(1):57-68

Metopograpsus: 4(1):112

Metridium senile Linné, 1758:
5(2):287-292

Miamira sinuata (van Hassett):
5(2):197-214

Micragenia Berry, 1953: 3(1):63-82

Micragenia oxystoma Berry, 1953:
3(1):63-82

Micrarionta (Eremarionta) aetotis Berry,
1928: 3(1):63-82

Micrarionta (Eremarionta) avawatzica
Berry, 1930: 3(1):63-82

Micrarionta (Eremarionta) borregonensis
Berry, 1929: 3(1):63-82

Micrarionta (Eremarionta) callinepius
Berry, 1930: 3(1):63-82

Micrarionta (Eremarionta) depressispira
Berry, 1928: 3(1):63-82

Micrarionta (Eremarionta) inglesiana
Berry, 1928: 3(1):63-82

Micrarionta (Eremarionta) melanopylon
Berry, 1930: 3(1):63-82

Micrarionta (Eremarionta) micrometalleus
Berry, 1930: 3(1):63-82

Micrarionta (Eremarionta) mille-palarum
Berry, 1930: 3(1):63-82

Micrarionta (Eremarionta) morongoana
Berry, 1930: 3(1):63-82

Micrarionta aquae-albae Berry, 1922:
3(1):63-82

Micrarionta opuntia Roth, 1975: 3(1):98;
4(2):237

Micrarionta sodalis (Hemphill, 1901):
3(1):98; 4(2):237

Micrarionta xerophila Berry, 1922:
3(1):63-82

Microciona astrosanguinea Bowerbank:
5(2):185-196

Micromelo: 5(2):185-196; $1:1-22

Micromelo undata (Bruguiere, 1792):
5(2):243-258

Micromenetus dilatatus (Gould): 3(1):99;
5(1):9-19

Micromya nebulosa (Conrad, 1834):
3(1):41-45

Micropogon undulatus (Linné): $3:59-70

Micropterus dolomieui (Lamarck): 5(1):1-7

Middendorffia caprearum (Sacchi):
6(1):57-68

Miesea Marcus, 1961: 5(2):183-184

Milax budapestensis (Hazy): 6(1):16

Milax gagates (Draparnaud, 1801): 6(1):16

Milax sowerbyi (Férussac): 6(1):16

Miliola marylandica Lea, 1833: 4(1):39-42

Minytrema melanops (Rafinesque):
$2:7-39, 89-94

Mistostigma Berry, 1947: 3(1):63-82

Mistostigma punctulum Berry, 1947:
3(1):63-82

Mitra idae Melville, 1893: 1:91-92

Mitra (Subcancilla) phorminx Berry, 1969:
3(1):63-82

Mitra (Tiara) caledinota Berry, 1960:
3(1):63-82

Mitra (Tiara) directa Berry, 1960: 3(1):63-82

Mitra (Tiara) lindsayi Berry, 1960: 3(1):63-82

Mitra montereyi Berry, 1920: 3(1):63-82

Mitra semiusta Berry, 1957: 3(1):63-82

Mitrella communis (Conrad, 1862):
4(1):39-42

Mitromica Berry, 1958: 3(1):63-82

Mitromorpha barbarensis woodfordi Berry,
1941: 3(1):63-82

Mitromorpha galeana Berry, 1941:
3(1):63-82

Mnemiopsis leidyi Agassiz: S3:59-70

Modiolus Lamarck, 1799: 1:108-109;
5(2):159-164 (passim); S1:23-24

Modiolus demissa Dillwyn, 1817: 3(1):33-40

Modiolus modiolus Linné, 1758: 3(1):33-40;
4(1):104; $1:59-78

Modulus Linne, 1758: 2:1-20

Mogula: 5(2):287-292 (passim)

Molgula manhattensis (DeKay): S3:59-70

(Mohavelix) Berry, 1943: 3(1):63-82

Mollusca, Unspecified: 2:79, 82, 84;
3(1):96-97, 107; 3(2):135-142 (passim);
4(1):115, 119, 119-120; 4(2):231,
238-239, 242

Monadenia: 3(1):3 (passim)

Monadenia (Corynadenia) tuolumneana
Berry, 1955: 3(1):63-82

Monadenia callipeplus Berry, 1940:
3(1):63-82 .

Monadenia chaceana Berry, 1940:
3(1):63-82

Monadenia cristulata Berry, 1940: 3(1):63-82

Monadenia fidelis: 2:98; 3(1):3 (passim)

Monadenia fidelis callidina Berry, 1940:
3(1):63-82

Monadenia fidelis celeuthia Berry, 1927:
3(1):63-82

Monadenia fidelis klamathica Berry, 1937:
3(1):63-82

Monadenia fidelis leonina Berry, 1937:
3(1):63-82

Monadenia fidelis ochromphalus Berry,
1937: 3(1):63-82

Monadenia fidelis pronotis Berry, 1931:
3(1):63-82

Monadenia fidelis scottiana Berry, 1940:
3(1):63-82

Monadenia fidelis smithiana Berry, 1940:
3(1):63-82

Monadenia infumata alamedensis Berry,
1940: 3(1):63-82

Monadenia marmarotis Berry, 1940:
3(1):63-82

Monadenia rotifer Berry, 1940: 3(1):63-82

Monas: S1:79-83

Moniliopsis chacei Berry, 1941: 3(1):63-82

Monochrysis lutheri (Droop): 3(1):33-40;
4(1):89-99; 6(2):189-197

Monodilepas Finley, 1927: 2:21-34

Monoplacophora ‘Wenz’ Knight, 1952:
$1:35-50

Mopalia Gray, 1847: 6(1):141-151

Mopalia chacei Berry, 1919: 3(1):63-82

Mopalia ciliata (Sowerby, 1840): 6(1):141-151

Mopalia cirrata Berry, 1919: 3(1):63-82

Mopalia cithara Berry, 1951: 3(1):63-82

Mopalia egretta Berry, 1919: 3(1):63-82

Mopalia hindsii (Reeve, 1847): 6(1):141-151

Mopalia lignosa (Gould, 1846):
6(1):141-151

Mopalia mucosa (Gould, 1846):
6(1):131-139, 141-151; S1:85-91

Mopalia phorminx Berry, 1919: 3(1):63-82

Mopalia (Dendrochiton) thamnopora
Berry, 1911: 3(1):63-82

Mopaliidae Dall, 1889: 6(1):141-151

Mordilla brockii Bergh, 1888: 5(2):243-258

Morone chrysops: S2:69-81

Moroteuthis pacifica: 4(2):241

Moroteuthis robusta: 4(2):241

Moschites adelieana Berry, 1917: 3(1):63-82

Moschites albida Berry, 1917: 3(1):63-82

Moschites aurorae Berry, 1917: 3(1):63-82

Moschites challengeri Berry, 1916:
3(1):63-82

Moschites harrissoni Berry, 1917: 3(1):63-82

Mourgona germaineae Marcus and Mar-
cus: 5(2):259-280

Mudéalia sp.: 1:27

Mulinia sp.: 4(1):104

Mulinia lateralis (Say, 1822): 2:35-40;
4(1):39-42; $1:59-78

Murex (Murex) tricornis Berry, 1960:
3(1):63-82

Murex acanthostephes Watson, 1883:
3(1):11-26

Murex carpenteri alba Berry, 1908:
3(1):63-82

Murex fulvescens Sowerby, 1834:
4(1):185-199 (passim)

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 243

Murex ramosus Linne, 1758: 4(1):109-110

Murex scala Gmelin, 1791: 2:57-61

Murex scabriculus Linné, 1758: 2:57-61

Murex semilunaris (Gmelin, 1791): 2:57-61

Muricanthus callidnus Berry, 1958:
3(1):63-82

Muricanthus nigritus (Philippi, 1845):
6(1):45-48

Muriciacea: 3(1):11-26

Muricidae: 3(1):11-26; 4(1):109-110; $1:1-22

Muricopsinae: 3(1):11-26

Musculista: 5(2):159-164 (passim)

Musculium Link, 1807: 3(2):269-272

Musculium lacustre (Miller, 1774):
3(2):187-200; 5(1):91-99

Musculium partumeium (Say, 1822):
3(2):187-200, 201-212; 5(1):49-64
(passim); S2:7-39, 193-201 (passim),
223-229

Musculium securis (Prime, 1861):
3(2):187-200; 5(1):21-30 (passim),
31-39, 49-64; $2:223-229

Musculium transverum (Say, 1829):
$2:223-229

Musculus: $1:23-34

Mya Linneé, 1758: 2:96

Mya arenaria Linné, 1758: 4(1):120-121;
6(2):179-188; S1:59-78, 79-83;
$3:59-70

Mya truncata Linne , 1758: 2:94;
4(1):120-121

Myochamidae Bronn, 1862: S1:35-50

Myrakeena: 4(2):157-162

Myrakeena angelica (Rochebrune, 1895):
4(2):157-162

Myrakeenini: 4(2):157-162

Myrina: $1:23-34

Mysella tumida (Carpenter, 1864):
4(2):234

Mytilacea: 2:41-50

Mytilidae Rafinesque, 1815: 1:101; 3(1):95;
$1:23-34

Mytilimeria nutalli Conrad: 5(1):173-176
(passim); S1:35-50

Mytilopsis leucophaeta (Conrad):
5(1):91-99 (passim)

Mytilopsis sallei (Recluz): 5(1):91-99
(passim)

Mytilus Linné, 1758: 1:108-109;
4(2):157-162; 5(1):41-48; 5(2):159-164;
$2:1-5 (passim)

Mytilus californianus Conrad, 1837:
3(1):33-40; S1:59-78

Mytilus canoasensis vidali ‘Ferreira and
Cuhna’ Woodring, 1973: 4(1):1-12

Mytilus desolationis: 1:105-106

Mytilus edulis Linné, 1758: 1:105-106, 108;
2:41-50, 63-73; 3(1):33-40; 3(2):179-186
(passim), 213-221; 4(1):104; 5(1):91-99
(passim); S1:35-50, 59-78, 79-83, 85-91

Mytilus galloprovincialis Lamarck, 1819:
1:105-106, 108; 5(1):91-99 (passim)

Myxa: $1:1-22

Nanostrea: 4(2):157-162

Nanostrea exigua Harry, 1985: 4(2):157-162

Nassa perpinquis bifasciata Berry, 1908:
3(1):63-82

Nassariidea Iredale, 1916: 3(1):101-102

Nassarius Dumeril, 1805: 2:57-71;
6(1):9-17

Nassarius obsoleta (Say, 1822):
4(2):165-172; 6(1):17

Nassarius pauperatus MckKillip and
Butler: 5(2):293-301 (passim)

Nassarius (Schizopyga) rhinetes Berry,
1953: 3(1):63-82

Nassarius trivittatus (Say, 1822):
4(2):165-172

Nassarius versicolor C. B. Adams, 1852:
4(1):1-12

Nautilus Linné, 1758: 4(2):217-227,
239-240; 6(1):69-78; S1:51-58

Nautilus macromphalus Sowerby, 1848:
2:90; S1:93-100

Nautilus pompilius Linné, 1758: 4(2):241

Navanax inermis (Cooper, 1863): 1:13
(passim); 5(2):287-292

Neda Mulsant, 1851: 5(2):215-241

Nekewis Stewart, 1927: 4(2):236

Nematolampas Berry, 1913: 3(1):63-82

Nematolampas regalis Berry, 1913:
3(1):63-82

Nematomenia banyulensis (Pruvot-Fol,
1951): 6(1):57-68

Nematomenia protecta (Odhner, 19347):
6(1):57-68

Nembrotha lineolata Bergh, 1905: 5(2):
243-258

Nembrotha livingstonei Allan, 1933:
5(2):243-258

Nemertea: 3(2):213-221

Neocorbicula Fischer, 1887: 5(2):243-258

Neogastropoda Wenz, 1941: S1:1-22, 23-34

Neoloricata Bergenhayn, 1955: 6(1):115-130

Neomenia Tullberg, 1878: S1:23-34

Neomenia carinata Tullberg, 1875:
6(1):57-68

Neomeniomorpha ‘Pelseneer’ Lankester,
1906: 5(2):281-286; 6(1):57-68

Neomphalace: S1:23-34

Neomphalidae: $1:23-34

Neomphalus fretterae McLean, 1981:
$1:23-34

Neopanope sayi (Smith): S3:59-70

Neopilina Lemche, 1957: 3(2):213-221;
6(1):57-68

Neopisidium Odhner, 1921: S2:223-229

Neopycnodonte Stenzel, 1971:
4(2):157-162

Neopycnodonte cochlear (Poli, 1795):
4(2):157-162

Neopycnodontini: 4(2):157-162

Neosimnia bella-maris Berry, 1946:
3(1):63-82

Neosimnia catalinensis Berry, 1916:
3(1):63-82

Neosimnia vidleri tyrianthina Berry, 1960:
3(1):63-82

Neothauma tanganyicense Smith, 1880:
4(1):107

Neotrigonia sp.: 4(1):13-19

Nereis: 2:96

Nerita clenchi Russell, 1940: 4(1):185-199
(passim)

Nerita forskali: 4(1):109-110

Nerita fulgurans: 3(2):223-231

Nerita funiculata Menke, 1852: 4(1):1-12

Nerita peloronta Linné, 1758: 4(1):185-199
(passim)

Neritacea Lamarck, 1816: 3(2):223-231

Neritidae Lamarck, 1816: 3(2):223-231;
4(1):109-110

Neritina latissima: 3(2):223-231

Neritina reclivata (Say, 1822): 4(1):185-199
(passim)

Neritina virginea Linné, 1758):
4(1):185-199 (passim)

Neverita (Glossaulax) andersoni (Clark,
1918): 4(1):1-12

Nitesselata Gmelin, 1791: 4(1):185-199
(passim)

Nitocris: S2:69-81

Nitzschia actinastroides (Lamm) von
Goor: 3(2):151-168

Nocomis micropogon (Cope): 5(1):1-7

Noetia ponderosa (Say, 1822): 4(1):111

Nomaeopelta Berry, 1958: 3(1):63-82

Nomaeopelta myrae Berry, 1959: 3(1):63-82

Notarchidae Eales, 1925: 5(2):243-258

Notaspidea Fischer, 1883: 5(2):215-241,
243-258; S1:1-22

Notobryon wardi Odhner, 1936:
5(2):243-258

Notoplax H. Adams, 1861: 6(1):115-130

Notoplax alisonae (‘Winckworth’ Kaas,
1976): 6(1):115-130

Notoplax coarcata (Sowerby, 1841):
6(1):115-130

Notoplax elegans Leloup, 1981:
6(1):115-130

Notoplax floridanus Dall, 1889: 6(1):79-114

Notoplax (Notoplax) arabica Kaas and
Van Belle, 1988, sp. nov.:
6(1):127-128

Notropis coccogenis (Cope): 5(1):1-7

Notropis galacturus (Cope): 5(1):1-7

Notropis spilopterus: S2:69-81

Noumea decussata Risbec, 1928:
5(2):243-258

Noumea purpurea Baba, 1949:
5(2):243-258

Noumea varians (Pease, 1871):
5(2):243-258

Nucella Roding, 1798: 4(1):110

Nucella emarginata (Deshayes, 1839):
1:105; 5(1):105-124 (passim)

Nucella lapillus (Linné, 1758): 1:92;
2:63-73; 4(1):110; 4(2):165-172;
5(1):105-124 (passim); S1:35-50

244 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Nucella lamellosa (Gmelin, 1791): 3(1):11-26

Nucinellidae: 4(1):111-112

Nucula (Ennucula) microsperma Berry,
1947: 3(1):63-82

Nucula sulcata (Bronn, 1831): 1:16 (passim)

Nuculacea Gray, 1824: 4(1):111-112

Nuculanacea H. and A. Adams, 1858:
A(1):111-112

Nudibranchia Cuvier, 1817: 2:84;
5(2):243-258, 281-286, 287-292;
$1:1-22

Nuphar luteum: 3(1):100

Nuttalina crossota Berry, 1956: 3(1):63-82

Nymphophilus minckleyi Taylor: 6(1):16

Obelia: 5(2):185-196, 287-292 (passim)

Obliquaria reflexa Rafinesque, 1820: 1:29,
43-50, 51-60; 2:85-86; 3(1):105;
4(1):25-37; 6(1):19-37

Obovaria Rafinesque, 1819: 4(1):117-118;
4(2):230-231

Obovaria jacksoniana (Frierson, 1912):
6(1):19-37

Obovaria olivaria (Rafinesque, 1820):
1:51-60; 3(1):105; 6(1):19-37

Obovaria retusa (Lamarck, 1819): 1:29,
31-34; 4(1):25-37; 6(1):19-37

Obovaria subrotunda (Rafinesque, 1820):
1:29, 31-34, 43-50; 2:85-86; 3(1):105;
4(1):21-23; 6(1):19-37; 6(2):165-178

Obovaria subrotunda lens (Lea, 1831):
1:29, 43-50; 4(1):25-37; 6(1):19-37

Obovaria subrotunda levigata (Rafinesque,
1820): 6(1):19-37; 6(2):165-178

Oceanebra crispatissima Berry, 1953:
3(1):63-82

Oceanebridae: 3(1):11-26

Octopodidae Rafinesque, 1815:
4(2):217-227; S1:93-100

Octopodinae: 2:89

Octopodoteuthidae Berry, 1912: 3(1):63-82

Octopoteuthidae Berry, 1912: 3(1):63-82

Octopus spp.: 2:89; 4(2):217-227, 233-234

Octopus alecto Berry, 1953: 3(1):63-82

Octopus bimaculoides Pickford and
McConnaughey, 1949: 2:90, 92;93,
93-94; 4(2):241-242

Octopus briareus Robson, 1929: 2:93-94;
4(2):217-227; 6(1):45-48

Octopus burryi Voss, 1950: 2:92;
6(2):207-211

Octopus defilippi Verany: 6(2):207-211

Octopus digueti Perrier and Rochebrune:
6(1):45-48; 6(2):207-211

Octopus dofleini (Wiilker, 1910): 2:90, 91;
4(2):241; 6(1):45-48; 6(2):207-211

Octopus dofleini martini Pickford 1964:
4(2):241

Octopus filosus Howell: 6(2):207-211

Octopus fitchi Berry: 1953: 3(1):63-82

Octopus hubbsorum Berry, 1953:
3(1):63-82

Octopus hummelincki Adam, 1936:
6(2):207-211

Octopus joubini Robson, 1929: 2:93-94;
6(1):45-48

Octopus maya Voss and Soliz Ramirez,
1966: 2:92, 93-94

Octopus micropyrsus Berry, 1953:
3(1):63-82

Octopus penicillifer Berry, 1954:
3(1):63-82

Octopus rubescens Berry, 1953:
3(1):63-82; 4(2):241

Octopus selene Voss, 1971: 6(2):207-211

Octopus tetricus Gould, 1852: 6(1):45-48

Octopus veligero Berry, 1953: 3(1):63-82

Octopus vulgaris Cuvier, 1797: 2:92;
4(2):217-227, 240; 6(1):45-48;
$1:35-50, 93-100

Ocythoe: 3(1):59 (passim); 4(2):217-227

Odontocymbiolinae: 3(1):11-26

Odostomia Fleming, 1813: S1:1-22;
$3:59-70

Odostomia (Chesapeakella): 3(1):96

Odostomia (Chlysallida): 4(1):122

Odostomia impressa (Say, 1821): 3(1):97

Oenopopota fidicula (Gould, 1849): 2:94-95

Oenopota levidensis (Carpenter, 1864):
2:94-95

Oenopota pumilus (Lea, 1833): 4(1):39-42

Oenopota turrispira Berry, 1941: 3(1):63-82

Offadesma angasi (Crosse and Fischer,
1864): 2:35-40

Ofina otis: 3(1):27-32 (passim)

Okadaia elegans Baba, 1930:
5(2):197-214, 243-258

Okenia mediterranea (Ihering, 1886):
5(2):243-258

Olea hansineensis Agersborg: 5(2):197-214

Oligochiton Berry, 1922: 3(1):63-82

Oligochiton lioplax Berry, 1922: 3(1):63-82

Oliva ionopsis Berry, 1969: 3(1):63-82

Olivella (Dactylidella) cymatilis Berry,
1963: 3(1):63-82

Olivella (Margintiella) walkeri Berry, 1958:
3(1):63-82

Olivella (Olivella) fletcherae Berry, 1958:
3(1):63-82

Olivella pynca Berry, 1935: 3(1):63-82

Omalogyra: $1:1-22

Ombrella Blainvile, 1824: 5(2):215-241

Ommastrephes bartrami: 2:89-90;
4(2):241

Ommastrephes hawaiiensis Berry, 1912:
3(1):63-82

Ommastrephoidea Berry, 1920: 3(1):63-82

Onchidella: $1:1-22

Onchidia: 2:21-34

Onchidiidae: 2:21-34; $1:1-22

Onchidium: $1:1-22

Onchidium verruculatum: 1:13 (passim)

Onchidorididae Gray, 1854: 5(2):243-258

Onchidoris aspera (Linne): 5(2):293-301

Onchidoris bilamellata (Linne):
5(2):197-214, 287-292, 293-301

Onchidoris hystricina (Bergh, 1878): 2:95

Onchidoris muricata (Miller, 1776): 2:95;
4(1):103-104; 5(2):197-214, 293-301

Onchidoris neapolitana (Delle Chiaje):
5(2):197-214

Onchidoris varians (Bergh, 1878): 2:95

Onchomelania hupensis: 2:88

Oncorhynchus kisutch (Walbaum):
5(2):125-128 (passim)

Oncorhynchus tshawytscha (Walbaum):
5(2):125-128 (passim)

Ondatra zibethica Linné, 1766:
6(2):165-178 (passim), 179-188
(passim)

Onithochiton Gray, 1847: 6(1):115-130

Onithochiton lyelli erythraeus Thiele, 1910:
6(1):115-130

Onithochiton erythraeus Thiele, 1910:
6(1):115-130

Onithochiton maillardi (Deshayes, 1863):
6(1):115-130

Onithochiton neglectus Rochebrune,
1881: 6(1):115-130

Onithochiton quercinus (Gould, 1846):
6(1):115-130

Onithochiton rugulosus Angas, 1867:
6(1):115-130

Onithochiton scholvieni Thiele, 1910:
6(1):115-130

Onithochiton titteratus (Krauss, 1848):
6(1):115-130

Onithochiton undilatus Quoy and
Gaimard, 1835: 6(1):115-130

Onithochiton wahibergi (Krauss, 1848):
6(1):115-130

Onoba: 4(1):185-199 (passim)

Onychoteuthis borealijaponica: 2:89-90

Opeatostoma Berry, 1958: 3(1):63-82

Operculatum H. and A. Adams, 1841:
5(2):215-241

Opisthobranchia Milne Edwards, 1848:
2:95-96; 5(2):281-286; S1:1-22

Opisthoteuthis californiana Berry, 1949:
3(1):63-82

Opisthoteuthis persephone Berry, 1918:
3(1):63-82

Opisthoteuthis pluto Berry, 1918:
3(1):63-82

Oplitaspongia pennata Lambe:
5(2):185-196, 197-214

Opsanus tau (Linné): $3:59-70

Opuntia littoralis: 2:98

Orbicularia: 5(2):159-164 (passim)

Orconectes immunis: S2:211-218

Orconectes proppinquus (Girard):
5(1):73-84

Orconectes rusticus (Girard): 5(1):73-84

Orconectes virilis (Hagen): 5(1):73-84

Oreohelicidae: 1:97, 2:98

Oreohelix californica Berry, 1931:
3(1):63-82

Oreohelix cooperi apiarium Berry, 1919:
3(1):63-82

Oreohelix flammulifer Berry, 1932: 3(1):63-82

AMER. MALAC. BULL. TAXONOMIC INDEX

Oreohelix handi jaegeri Berry, 1931:
3(1):63-82

Oreohelix nevadensis Berry, 1932:
3(1):63-82

Oreohelix strigosa canadica Berry, 1932:
3(1):63-82

Oreohelix vortex Berry, 1932: 3(1):63-82

Orthalicus floridensis Pilsbry, 1891: 2:98

Orthalicus reses (Say): 2:98

Orthalicus reses nesodryas Pilsbry, 1946:
2:98

Orthalicus undulatus jamaicensis Pilsbry,
1899: 2:98

Orymaeus: 4(1):113-114

Oscaniopsis Bergh, 1897: 5(2):215-241

Oscaniella Bergh, 1897: 5(2):215-241

Oscanius Gray, 1847: 5(2):215-241

Ostrea Linné, 1758: 1:90; 4(1):1-12;
4(2):157-162

Ostrea chilensis Philippi: S3:1-4

Ostrea denselamellosa Lischke, 1869:
4(2):157-162

Ostrea edulis Linné, 1758: 1:105-106;
4(1):61-79 (passim); 4(2):157-162;
$1:35-50; S3:41-49

Ostrea (Eostrea) |hering, 1907:
4(2):157-162

Ostrea (Eostrea) puelchana Orbigny,
1846: 4(2):157-162

Ostrea equestris Say, 1834: 2:63-73

Ostrea gigas Thunberg, 1793: 3(1):85-88

Ostrea irridescens Hanley, 1854: 4(1):119

Ostrea lurida Carpenter, 1864: 1:102;
4(1):61-79 (passim)

Ostreidae Rafinesque, 1815: 2:41-50;
4(2):157-162

Ostreinae: 4(2):157-162

Ostreini: 4(2):157-162

Ostreola: 4(2):157-162

Ostreola conchaphila (Carpenter, 1857):
4(2):157-162

Ostreola equestris (Say, 1834):
4(2):157-162

Ostreola stentina (Payraudeau, 1826):
4(2):157-162

Otala lactea Miiller: 6(1):16

Otina: $1:1-22

Otinidae: $1:1-22

Ovatella: $1:1-22

Oxychilus cellarius (Miller, 1774): 6(1):16

Oxynidae: $1:1-22

Oxynoe: $1:1-22

Oxynoe antillarum Fischer: 5(2):259-280

Oxynoe azuropunctata Jensen:
5(2):197-214, 259-280

Oxynoe viridis (Pease, 1861): 5(2):243-258

Oxynoidae: 5(2):243-258

Pachythaerus: 4(2):238

Pachygrapsus crassipes: 2:1-20

(Pagodula) Monterosato, 1884:
3(1):101-102

Paleoheterodonta Newell, 1965:
4(1):111-112

Pallifera: 4(2):238

Palythoa: 5(2):185-186

Panacca africana Fischer: 3(1):103-104

Panacca arata Verrill and Smith, 1881:
3(1):103-104

Panacca fragilis Grieg: 3(1):103-104

Panacca locardi Dall, 1903: 3(1):103-104

Pandoracea Rafinesque, 1815: 2:35-40

Pandoridae Rafinesque, 1815: S1:35-50

Panopeus herbstii (Milne-Edwards):
2:1-20; S3:59-70

Paraganitus ellynnae Challis: 5(2):281-286

Parahyotissa: 4(2):157-162

Parahyotissa imbricata (Lamarck, 1819):
4(2):157-162

Parahyotissa mcgintyi Harry, 1985:
4(2):157-162

Parahyotissa (Numismoida): 4(2):157-162

Parahyotissa (Numismoida) numisma
(Lamarck, 1819): 4(2):157-162

Parahyotissa (Pliohyotissa): 4(2):157-162

Parahyotissa (Pliohyotissa) quercinus
(Sowerby, 1819): 4(2):157-162

Paralabrax maculatofasciatus Stein-
dacher: 6(1):45-48

Parilimya fragilis (Gould): $1:35-50

Parilimyidae Morton, 1981: S1:35-50

Parmophorus Cantraine, 1835:
5(2):215-241

Partula: 6(1):9-17

Partula gibba Bruguiere: 6(1):16

Partula mirabilis Crampton: 6(1):16

Partula mooreana: 1:103-104

Partula olympia Crampton: 6(1):16

Partula otaheitana Ferussac: 6(1):16

Partula suturalis Pfeiffer: 1:103-104; 6(1):16

Partula taeniata Morch: 1:103-104; 6(1):16

Patella Linné, 1758: 4(1):115

Patella aspersa: 3(1):33-40

Patella perversa Gmelin, 1790:
5(2):215-241

Patella umbraculum Lightfoot, 1786:
5(2):215-241

Patella vulgata: 3(1):33-40; 3(2):223-231;
$1:35-50

Patellidae: 3(1):95; 4(1):115

Patellogastropoda: 4(1):115

Paziella: 3(1):11-26

Paziella pazi (Crosse, 1869): 3(1):11-26

Pecten Miller, 1776: 1:13 (passim);
4(2):157-162

Pecten (Leptopecten) euterpes Berry,
1957: 3(1):63-82

Pecten lunaris Berry, 1963: 3(1):63-82

Pecten maximus (Linné, 1758): $1:35-50

Pectinacea Rafinesque, 1815: 2:41-50;
4(1):111-112

Pedicularia (californica?) ovuliformis Berry,
1946: 3(1):63-82

Pegias Simpson, 1900: 4(1):117-118

Pegias fabula (Lea, 1836): 1:43-50;
6(1):19-37

Pegmapex Berry, 1960: 3(1):63-82

: 1983 - 1988 245

Pegmapex phoebe Berry, 1960: 3(1):63-82

Pelecypoda, Unspecified: 2:79

Peloscolex ferox: S2:7-39

Pelseneeria spp.: 2:83

Peltodoris atromaculata Bergh: 4(2):232;
5(2):185-196, 197-214

Penicillus dumetosus (Lamouroux) Blain-
ville: 5(2):259-280

Peracle: $1:1-22

Peraclidae: S1:1-22

Periplaneta americana: S1:79-83

Periploma fragile Totten, 1835: 2:35-40;
$1:35-50

Periploma margaritaceum (Lamarck,
1801): 2:35-40

Periploma (Offadesma) angasi Crosse
and Fischer: S1:35-50

Periploma orbiculare Guppy, 1882: 2:35-40

Periploma ovata: 2:35-40

Periplomatidae: 2:35-40; S1:35-50

Perissitys Stewart, 1927: 4(2):236

Perkinsus marinus (Mackin, Owen and
Collier): S3:59-70

Perna canaliculus (Gmelin): 5(2):159-164
(passim)

Perna perna: 5(2):159-164 (passim)

Perna viridis (Linné, 1758): 4(2):233;
5(2):159-164

Persicula pulchella (Kiener, 1834): 2:84

Petromyzon marinus (Linné): 5(1):21-30
(passim)

Phaenommia Morch, 1860: 2:1-20

Phanerophthalmus: $1:1-22

Phanerophthalmus smaragdius (Ruippell
and Leuckhart, 1831): 5(2):243-258

Phascolosoma agassizii: 1:91-92

Phestilla: 5(2):287-292

Phestilla lugubris Bergh: 5(2):185-186

Phestilla melanobranchia Bergh, 1874:
5(2):185-196, 197-214, 243-258

Phestilla minor Rudman: 5(2):185-196

Phestilla sibogae Bergh: 5(2):185-196,
197-214, 293-301 (passim)

Phidiana crassicornis (Eschscholtz):
5(2):197-214

Philinacea: 4(2):233

Philine: $1:1-22

Philine angasi Crosse and Fischer:
5(2):185-196

Philine aperta (Linné): 5(2):185-196

Philine auriformis Suter: 5(2):185-196

Philine gibba Strebel: 5(2):197-214

Philine lima (Brown): 5(2):185-196

Philine scabra (Miller): 5(2):185-196

Philine thurmanni Marcus and Marcus:
5(2):185-196

Philinidae: $1:1-22

Philinoglossa: 5(2):281-286; S1:1-22

Philinoglossa marcusi Challis, 1969:
5(2):281-286

Philinoglossidae: $1:1-22

Philinopsis capensis (Bergh, 1907):
5(2):243-258

246 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Philinopsis cyanea (Martens, 1879):
5(2):243-258

Philippia: S1:1-22

Philippia (Basisulcata) Melone and
Taviana, 1985: 4(1):108-109

Philippia (Philippia) Gray, 1847: 4(1):108-109

Philippia (Psilaxis) Woodring, 1928:
4(1):108-109

Pholadidae: S1:59-78

Pholadomya candida Sowerby: S1:35-50

Pholadomyidae Gray, 1947: S1:35-50

Pogonias cromis (Linne): $3:59-70

Phyllaplysia engeli Marcus: 5(2):197-214

Phyllaplysia taylori: 4(2):205-216 (passim);
5(2):197-214

Phyllaplysia zostericola McCauley:
5(2):185-196

Phyllida: 5(2):243-258

Phyllida varicosa Lamarck, 1801:
4(1):109-110; 5(2):185-196, 243-258

Phyllidiidae: 5(2):243-258

Phylliroe bucephala Peron and Lesueur:
5(2):197-214

Phyllobranchillus orientalis: 4(1):109-111

Phyllodesmium cryptica Rudman:
5(2):185-196

Phyllodesmium hyalinum Ehrenberg, 1931:
5(2):185-196, 243-258

Phyllodesmium poindimiei (Risbec, 1928):
5(2):185-196, 243-258

Phyllodesmium serratum (Baba, 1949):
5(2):243-258

Phyllodesmium xeniae: 4(1):109-111

Phylomycidae: 4(2):238

Phylomycus carolinianus: 4(2):238

Phylomycus togatus: 4(2):238

Physa sp.: 1:31-34; 6(1):57-68; S2:69-81

Physa ancillaria Say, 1825: 5(1):9-19

Physa fontinalis (Linné, 1758): 3(2):135-142
(passim), 243-265; 5(1):65-72 (passim)

Physa heterostropha (Say, 1817): 5(1):9-19;
6(1):17

Physa integra Haldeman, 1841: 5(1):73-84

Physa propinqua Tyron, 1865: 5(1):65-72
(passim)

Physella ancellaria (Say, 1825): 3(1):99

Physella gyrina (Say, 1821): 5(1):31-39,
105-124 (passim); 6(2):165-178

Physella integra (Haldeman, 1841):
5(1):105-124 (passim)

Physella virgata (Gould, 1855): 3(2):269-272

Physella virgata virgata (Gould, 1855):
3(2):243-265

Pilina: 6(1):69-78

Pinctada martensi (Dunker, 1868): 1:101;
5(1):173-176 (passim)

Pinctada mazatlanica (Hanley, 1856):
4(1):119

Pinna muricata Linné, 1758: 6(1):115-130

Pinna pectinata Linné, 1767: 4(2):217-227

Pinnidae Leach, 1819: 2:97

Pinufius rebus Marcus and Marcus, 1960:
5(2):185-196

Pisania maculosa: 3(1):27-32 (passim)

Pisaster ochraceus (Brandt, 1835):
6(1):141-151

Piseinotecus Marcus, 1961: 5(2):183-184

Piseinotecus sphaeriferus (Schmekel,
1965): 5(2):197-214

Pisidiidae Gray, 1857: 3(1):100;
3(2):201-212; 4(1):61-79, 116

Pisidium Pfeiffer, 1821: 3(2):269-272;
$2:187-191, 193-201 (passim)

Pisidium amnicum (Miller, 1774):
3(2):187-200; 5(1):21-30 (passim), 41-48

Pisidium annandalei Prashad: 5(1):91-99

Pisidium casertanum (Poli, 1795):
3(2):187-200, 201-212; 4(1):116;
5(1):1-7, 21-30, 31-39, 49-64;
$2:223-229

Pisidium clarkeanum G. and H. Nevill:
5(1):91-99

Pisidium compressum Prime, 1852:
3(2):187-200; 5(1):1-7, 31-39;
$2:223-229

Pisidium conventus Clessin, 1877:
3(2):187-200; 5(1):21-30; S2:219-222,
223-229

Pisidium crassum Sterki, 1901:
3(2):187-200

Pisidium dubium (Say, 1816): 3(2):187-200

Pisidium equilaterale Prime, 1852:
$2:223-229

Pisidium ferrugineum Prime, 1852:
3(2):187-200; 5(1):31-39

Pisidium henslowanum (Sheppard, 1825):
3(2):187-200

Pisidium lilljeborgi Clessin, 1886:
3(2):187-200

Pisidium moitessierianum Paladilhe, 1866:
5(1):21-30 (passim)

Pisidium nitidum Held, 1836: 3(2):187-200

Pisidium obtusale Pfeiffer, 1821:
3(2):187-200

Pisidium personatum Malm, 1855:
3(2):187-200; 5(1):41-48

Pisidium punctatum Sterki, 1895:
$2:223-229

Pisidium subtruncatum Malam, 1855:
3(2):187-200

Pisidium ultramontanum Prime, 1865:
$2:223-229

Pisidium variable Prime, 1852:
3(2):187-200; 5(1):31-39; S2:223-229

Pisidium ventricosus Prime, 1851:
3(2):187-200

Pisidium walkeri Sterki, 1895: 3(2):187-200;
$2:223-229

Pitar (Lamelliconcha) hesperius Berry,
1960: 3(1):63-82

Placida cremoniana (Trichese):
5(2):197-214

Placida dendritica (Alder and Hancock,
1843): 5(2):243-258, 259-280

Placida kingstoni (Thompson):
5(2):259-280

Placida viridis (Trinchese): 5(2):197-214

Placiphorella ‘Carpenter’ Dall, 1879:
6(1):141-151

Placiphorella pacifica Berry, 1919:
3(1):63-82

Placiphorella rufa Berry, 1917: 3(1):63-82

Placiphorella stimpsoni (Gould, 1859):
6(1):141-151

Placiphorella velata Dall, 1878:
6(1):141-151

Placopecten magellanicus (Gmelin, 1791):
4(1):104; 6(1):1-8; S1:59-78

Placuna ‘Solander’ Lightfoot, 1786:
4(2):157-162 (passim)

Plagiola interrupta (Rafinesque, 1820):
6(1):19-37

Plagiola lineolata (Say, 1834): 1:29, 43-50

Plagiola lineolata (Rafinesque, 1820):
6(1):19-37

Plagioporus hypentelli Hendrix, 1973:
4(1):119

Planaxidae Gray, 1850: 3(1):96; 4(2):235

Planaxis Lamarck, 1822: 2:1-20; 3(1):96

Planorbidae Gray, 1840: $1:1-22

Planorbis corneus (Linné, 1758):
3(2):135-142, 213-221; 5(1):105-124
(passim)

Planorbis planorbis (Linné, 1758): 5(1):65-72

Planorbis vortex (Linné, 1758): 5(1):65-72,
73-84 (passim)

Planorbula armigera (Say, 1818): 3(1):99;
5(1):9-19

Planostrea: 4(2):157-162

Planostrea pestigris (Hanley, 1846):
4(2):157-162

Platydorididae: 5(2):243-258

Platydoris cruenta (Quoy and Gaimard,
1832): 5(2):243-258

Platydoris scabra (Cuvier, 1806):
5(2):197-214, 243-258

Plaxiphora obtecta (‘Carpenter’ Pilsbry,
1893): 6(1):141-151

Plectomerus dombeyanus (Valenciennes,
1833): 6(1):19-37

Pleioptygma Conrad, 1863: 3(1):97-98

Pleioptygma helenae Radwin and Bibbey,
1972: 3(1):97-98

Plethobasus Simpson, 1900: 6(2):165-178

Plethobasus cicatricosus (Say, 1829):
4(1):25-37; 6(1):19-37

Plethobasus cooperianus: 4(1):25-37;
6(1):19-37, 49-54; 6(2):165-178

Plethobasus cyphyus (Rafinesque, 1820):
1:29, 51-60; 2:85-86; 4(1):25-37;
5(2):165-171; 6(1):19-37; 6(2):165-178

Plethobasus cyphyus compterus (Frier-
son, 1911): 6(1):19-37

Plethobasus pachosteus (Rafinesque,
1820): 6(1):19-37

Plethobasus striatus (Rafinesque, 1820):
4(1):25-37; 6(1):19-37

Pleurehdera Marcus and Marcus, 1970:
5(2):215-241

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 247

Pleurehdera haraldi (Marcus and Marcus,
1970): 5(2):215-241

Pleurobema Rafinesque, 1820: 6(2):165-178

Pleurobema aldrichianum Goodrich, 1931:
6(1):19-37

Pleurobema clava (Lamarck, 1819):
1:31-34; 4(1):25-37; 6(1):19-37;
6(2):165-178

Pleurobema clava catillus (Conrad, 1836):
6(1):19-37

Pleurobema coccineum (Conrad, 1836):
1:29; 2:85; 3(1):105; 6(1):19-37

Pleurobema cordatum (Rafinesque, 1820):
1:29, 31-34, 43-50; 2:85-86;
4(1):25-37; 6(1):19-37; 6(2):165-178

Pleurobema gibberum (Lea, 1838):
6(1):19-37

Pleurobema obliguum Lamarck, 1819:
6(1):19-37

Pleurobema obliquata Rafinesque, 1820:
6(1):19-37

Pleurobema obliqum (Lamarck, 1819):
3(1):41-44; 4(1):25-37

Pleurobema oviforme (Conrad, 1834):
1:43-50; 3(1):41-44, 104, 106; 5(1):1-7;
6(1):19-37; 6(2):165-178, 179-188

Pleurobema oviforme argenteum (Lea,
1841): 3(1):41-44; 6(1):19-37;
6(2):165-178

Pleurobema oviforme holstonense (Lea,
1840): 6(1):19-37

Pleurobema permorsa Rafinesque, 1831:
6(1):19-37

Pleurobema plenum (Lea, 1840): 1:28, 29,
31-34, 43-50; 4(1):25-37, 117;
6(1):19-37; 6(2):165-178

Pleurobema pyramidatum (Lea, 1834):
1:29; 4(1):25-37; 6(1):19-37

Pleurobema rubrum (Rafinesque, 1820):
1:31-34, 51-60; 2:85-86; 4(1):25-37;
6(1):19-37; 6(2):165-178

Pleurobema sintoxia (Rafinesque, 1820):
1:31-34, 51-60; 2:85-86

Pleurobranchacea Menke, 1828:
5(2):215-241

Pleurobranchaea ‘Meckel’ Leve, 1813:
5(2):215-241

Pleurobranchaea californica (Dall, 1900):
5(2):287-292

Pleurobranchaea maculata (Quoy and
Gaimard): 5(2):215-241

Pleurobranchaea meckelii Blainville, 1825:
§(2):215-241

Pleurobranchaeidae Pilsbry, 1896:
5(2):215-241, 243-258

Pleurobranchella Thiele, 1925: 5(2):215-241

Pleurobranchella alba (Guangyu and Si):
5(2):215-241

Pleurobranchella nicobarica Thiele:
5(2):215-241

Pleurobranchia: 5(2):185-196

Pleurobranchidae Menke, 1828:
§(2):215-241, 243-258; $1:1-22

Pleurobranchidium Blainville, 1825:
5(2):215-241

Pleurobranchillus Bergh, 1892:
5(2):215-241

Pleurobranchinae Ferussac, 1822:
5(2):215-241

Pleurobranchoides gilchristi O'Donoghue,
1929: 5(2):215-241

Pleurobranchomorpha: S1:1-22

Pleurobranchus Cuvier, 1805:
5(2):215-241; $1:1-22

Pleurobranchus albiguttatus Bergh:
5(2):215-241

Pleurobranchus brockii Bergh, 1897:
5(2):243-258

Pleurobranchus bubala Marcus and
Gosliner, 1984: 5(2):243-258

Pleurobranchus forsskali Ruppel and
Leuckart: 5(2):215-241

Pleurobranchus grandis Pease: 5(2):215-241

Pleurobranchus inhacae Macnae, 1962:
5(2):243-258

Pleurobranchus luniceps Cuvier, 1817:
5(2):215-241

Pleurobranchus mamillatus Quoy and
Gaimard: 5(2):215-241

Pleurobranchus membranceus: 5(2):215-241

Pleurobranchus nigropunctatus (Bergh,
1907): 5(2):243-258

Pleurobranchus ovalis: 5(2):215-241

Pleurobranchus peronii Cuvier, 1805:
5(2):215-241, 243-258

Pleurobranchus tarda Verrill, 1880:
5(2):243-258

Pleurobranchus xhosa Macnae, 1962:
5(2):243-258

Pleurocera acuta Rafinesque, 1831:
3(1):100

Pleurocera alvare (Conrad, 1834):
4(1):25-37

Pleurocera canaliculatum (Say, 1821):
1:31-34, 51-60; 4(1):25-37;
6(2):165-178

Pleurocera canaliculatum undulatum (Say,
1829): 4(1):25-37

Pleurocera parvum (Lea, 1862):
6(2):165-178

Pleuroceridae: 3(2):223-231

Pleurodonte: 3(1):102-103

Pleuroliria artia Berry, 1957: 3(1):63-82

Pleuroliria parthenia Berry, 1957: 3(1):63-82

Pleuroploca trapezuim (sic): 4(1):109-110

Pleurotomaria atlantica (Ricos and
Matthews, 1968): 3(1):101-102

Plicatula inezana Durham, 1950: 4(1):1-12

Pliodon Agassiz, 1846: 4(1):107

Pliodon ovata (Swainson, 1832): 4(1):107

Plidon spekii (Woodward, 1859): 4(1):107

Plocamopherus gulo: 5(2):183-184

Plocamopherus imperialis Angas, 1864:
5(2):185-196

Plocamopherus maculatus (Pease, 1860):
5(2):243-258

Ploiochiton Berry, 1926: 3(1):63-82

Pogonophora: S$1:23-34

Poirieri: 3(1):11-26

Polinices sp.: 4(1):185-199 (passim)

Polinices duplicatus (Say, 1822):
3(2):135-142 (passim); 4(1);111

Polita gabrielina Berry, 1924: 3(1):63-82

Polycelis tenuis: 3(2):213-221 (passim)

Polycera Cuvier, 1817: 6(1):57-68

Polycera capensis Quoy and Gaimard,
1824: 5(2):243-258

Polycera elegans (Bergh, 1869):
5(2):185-196

Polycera faeroensis Lemche, 1929:
5(2):185-196

Polycera hedgpethi Marcus, 1964:
5(2):243-258

Polycera quadrilineata (Miller, 1776):
5(2):185-196, 197-214, 243-258

Polycera zosterae O’Donoghue, 1924:
5(2):197-214

Polycera emertoni Verrill, 1880:
5(2):197-214

Polyceridae: 5(2):243-258

Polygyra columbiana oria Berry, 1933:
3(1):63-82

Polygyra columbiana shasta Berry, 1921:
3(1):63-82

Polygyra hapla Berry, 1933: 3(1):63-82

Polygyra loricata nortensis Berry, 1933:
3(1):63-82

Polygyra pinicola Berry, 1916: 3(1):63-82

Polygyra sierrana Berry, 1921: 3(1):63-82

Polygyra trachypepla Berry, 1933:
3(1):63-82

Polygyridae Pilsbry, 1930: 2:98

Polymesoda anomala (Deshayes, 1855):
6(2):199-206 (passim)

Polymesoda caroliniana Bosc, 1801:
4(1):116-117; 4(2):234; 6(2):199-206

Polymesoda (Geloina) erosa (Solander):
5(1):21-30 (passim), 91-99

Polymita: 3(1):102-103

Polyplacophora Blainville, 1816: 1:99;
6(1):57-68, 115-130; S1:35-50

Polypodoidea Berry, 1920: 3(1):63-82

Polypus (Pinnoctopus?) kermadecensis
Berry, 1914: 3(1):63-82

Polypus apollyon Berry, 1912: 3(1):63-82

Polypus californicus Berry, 1911:
3(1):63-82

Polypus gilbertianus Berry, 1912:
3(1):63-82

Polypus hokkaidoensis Berry, 1921:
3(1):63-82

Polypus hoylei Berry, 1909: 3(1):63-82

Polypus leioderma Berry, 1911: 3(1):63-82

Polypus madokai Berry, 1921: 3(1):63-82

Polypus oliveri Berry, 1914: 3(1):63-82

Polypus pricei Berry, 1913: 3(1):63-82

Polypus scorpio Berry, 1920: 3(1):63-82

Polystira barrettii (Guppy, 1866):
4(1):185-199 (passim)

248 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Pomacea lineata: 3(2):223-231

Pomacea paludosa: 1:97; S1:51-58

Pomatias elegans: 3(1):27-32 (passim)

Pontohedyle milaschewitschii (Kowalevsky,
1901): 5(2):303-306

Popenaias popei (Lea, 1857): 2:86

Porites somaliensis: 5(2):185-196, 197-214

Porpita: 5(2):185-196

Potamides obtusus: 2:1-20

Potamides quadratus Sowerby: 2:1-20

Potamides telescopium: 2:1-20

Potamididae H. and A. Adams, 1854: 2:1-20

Potamidinae H. and A. Adams, 1854:
2:1-20

Potamilus Rafinesque, 1818: 4(1):117-118

Potamilus alatus (Say, 1817): 1:51-60,
71-74; 2:85-86; 3(1):41-45, 47-53;
4(1):25-37, 117; 5(2):165-171;
6(1):19-37; 6(2):165-178, 179-188

Potamilus capax (Green, 1832):
4(2):230-231

Potamilus ohiensis (Rafinesque, 1820):
1:51-60, 71-74

Potamilus ohioensis (Rafinesque, 1820):
6(1):19-37

Potamilus purpurata (Lamarck, 1819):
4(1):21-23; 6(1):19-37

Potamogeton: 5(1):65-72 (passim), 73-84

Potamopyrgus jenkinsii (Smith):
3(2):223-231; 5(1):73-84; 6(1):17

Precuthona divae Marcus, 1861:
5(2):197-214

Prinocidaris hawaiiensis: 2:83

Procambarus Clarkii: S2:89-94, 211-218

Prochaetoderma: 6(1):57-68

Prochaetodermatidae: 3(1):97

Procladius culiciformis: S2:7-39

Procyon lotor: S2:7-39, 89-94

Profissurellidea Wenz, 1938: 2:21-34

Promenetus exacuous (Say): 3(1)99;
5(1):9-19, 73-84

Proptera alata (Say, 1817): 1:29, 43-50;
3(1):105; 6(1):19-37

Proptera laevissima (Lea, 1830): 6(1):19-37

Prosobranchia Milne Edwards, 1848:
$1:1-22, 23-34

Protobranchia Pelseneer, 1889: 4(1):111-112

Protostomia: 3(2):213-221 (passim)

Protothaca Dall, 1902: 4(1):1-12

Protothaca asperima (Sowerby, 1835):
4(1):119

Pruvottfoilia pselliotes (Labbé, 1923):
5(2):243-258

Psephidia brunnea Dall, 1916: 3(1):103

Pseudomalaxis Fischer, 1885: S1:1-22

Pseudomalaxis (Pseudomalaxis) Fischer,
1885: 4(1):108-109

Pseudomalaxis (Spirolaxis) Monterosato,
1913: 4(1):108-109

Pseudomelampus mexicanus Berry, 1964:
3(1):63-82

Pseudomelatoma sticta Berry, 1956:
3(1):63-82

Pseudomiltha Fischer, 1885: S1:23-34

Pseudomonas stutzeri: 2:93-94

Pseudopleuronectes americanus
(Walbaum): 5(2):287-292

Pseudoskenella: $1:1-22

Pseudosuccinea columella (Say, 1821):
3(1):99; 5(1):9-19; 6(2):165-178

Pseudovermis Periaslavzeff, 1891: 2:95;
5(2):281-286

Pseudovermis hancocki Challis:
5(2):281-286

Pseudovermis mortoni Challis: 5(2):281-286

Pseudunela: 5(2):281-286

Pseudunela cornuta (Challis):
5(2):281-286

Ptenoglossa Gray, 1853: $1:1-22

Pteraeolidia ianthina (Angas, 1864):
5(2):197-214

Pteroctopus tetracirrhus (Delle Chiaje,
1830): 4(2):217-227; 6(2):207-211

Pteropoda Cuvier, 1804: 5(2):185-196

Pteropurpura (Centrifuga) deroyana Berry,
1963: 3(1):63-82

Pterygioteuthis microlampas Berry, 1913:
3(1):63-82

Ptychobranchus Simpson, 1900:
4(1):117-118; 6(2):165-178

Ptychobranchus fasciolare (Rafinesque,
1820): 1:29; 3(1):105; 6(1):19-37

Ptychobranchus fasciolaris (Rafinesque,
1820): 1:29, 31-34, 43-50; 2:85-86;
3(1):41-45, 47-53, 104; 4(1):25-37;
6(2):165-178

Ptychobranchus occidentalis (Conrad,
1836): 2:85

Ptychobranchus subtentum (Say, 1825):
1:43-50; 3(1):41-45, 104; 4(1):25-37;
6(1):19-37; 6(2):165-178

Ptychosyrinx chilensis Berry, 1968:
3(1):63-82

Pulmonata Cuvier, 1817: S1:1-22

Puncturella punctocostata Berry, 1947:
3(1):63-82

Puncturella ralphi Berry, 1947: 3(1):63-82

Pupa Roding, 1798: S1:1-22

Pupa affinis (A. Adams, 1854):
5(2):243-258

Pupa kirki (Hutton): 5(2):185-196

Pupa solidula (Linné, 1758): 5(2):243-258

Pupa sulcata (Gmelin, 1791): 5(2):243-258

Pupa suturalis (A. Adams, 1854):
5(2):243-258

Pupa tessellata (Reeve, 1842): 5(2):243-258

Puperita pupa (Linne, 1767): 4(1):185-199

Pupilla blandi Morse, 1865: 1:99

Pupilla hebes (Ancey, 1881): 1:99

Pupilla muscorum (Linne, 1758): 1:99

Pupilla sonorana (Sterki, 1899): 1:99

Pupilla sterkiana (Pilsbry, 1889): 1:99

Pupilla syngenes (Pilsbry, 1890): 1:99

Pupillaea Sowerby, 1835: 2:21-34

Pupillaea annulus (Odhner, 1932): 2:21-34

Pupillaea aperta (Sowerby, 1825): 2:21-34

Pupillaea aperta teheulcha \hering, 1907:
2:21-34

Purisima: 2:84-85

Purpura Bruguiére, 1789: 3(1):101-102;
4(1):110

Purpura patula (Linné, 1758): $1:1-22

Purpura persica (Linné, 1758): 4(1):110

Purpurella Dall, 1871: 4(1):110

Purpurella patula (Linné, 1758): 4(1):110

Pustulostrea: 4(2):157-162

Pustulostrea tuberculata (Lamarck, 1804):
4(2):157-162

Pustulostrini: 4(2):157-162

Pycnodonte Fischer, 1835: 4(2):157-162

Pycodonteninae Stenzel, 1959:
4(2):157-162

Pycnopodia helianthoides: 5(2):185-196

Pyramidella crenulata (Holmes, 1859):
$1:1-22

Pyramidellacea Gray, 1847: S1:1-22

Pyramidellidae Gray, 1847: 3(1):96; S1:1-22

Pyrazus Montfort, 1910: 2:1-20

Pyrazus ebininus (Bruguiere, 1792): 2:1-20

Pyrgopsis lemur Berry, 1920: 3(1):63-82

Pyrgulopsis archimedis Berry, 1947:
3(1):63-82

Pythia Roding, 1798: S1:1-22

Quadrula Rafinesque, 1820: 1:109-110;
6(2):165-178, 179-188 (passim)

Quaadrula apiculata (Say, 1829): 2:86

Quaadrula bullata (Rafinesque, 1820):
6(1):19-37

Quadrula cylindrica (Say, 1817): 1:28,
43-50; 4(1):25-37; 6(1):19-37;
6(2):165-178

Quadtrula cylindrica cylindrica (Say, 1817):
4(1):117-118

Quadrula cylindrica strigillata (Wright,
1898): 6(1):19-37

Quaarula fragosa (Conrad, 1835):
4(2):230-231; 6(1):19-37

Quaarula intermedia (Conrad, 1836):
1:43-50; 3(1):41-45; 4(1):25-37;
6(1):19-37

Quadrula metanerva Rafinesque, 1820:
1:29, 43-50, 51-60; 4(1):25-37;
4(2):230-231; 5(2):165-171; 6(2):19-37

Quadrula nodulata (Rafinesque, 1820):
6(1):19-37

Quaadrula nodulata (Say, 1834): 1:29, 51-60

Quadrula pustulosa (Lea, 1831): 1:29, 31,
34, 43-50; 3(1):105; 4(1):21-23;
4(1)25-37; 5(2):165-171; 6(1):19-37;
6(2):165-178

Quadrula pustulosa pustulosa (Lea, 1831):
1:51-60; 2:85-86; 4(1):117-118

Quaarula quaarula (Rafinesque, 1820):
1:29, 31-34, 43-50, 51-60, 71-74;
3(1):105; 5(2):165-171; S2:101
(passim); 6(1):19-37

Quaarula sparsa (Lea, 1841): 3(1):41-45;
6(1):19-37; 6(2):165-178

Quibulla Iredale, 1929: 5(2):185-196

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 249

Quincuncina Ortmann, 1922: 1:109-110

Quincuncina infucata (Conrad, 1834):
$2:7-39

Rabdotus baileyi (Dall, 1893): 4(1):113-114

Rabdotus nigromontanus (Dall, 1897):
4(1):113-114

Rachiglossa Gray, 1853: 3(1):11-26

Radiocentrum avalonense Hemphill, 1902:
2:98

Radix: 2:88; S1:1-22

Radix limosa (Linné): 5(1):65-72 (passim)

Radix peregia: 3(1):27-32 (passim)

Radix quadrasi (Bequaert and Clench):
5(1):105-124 (passim)

Raeta: 4(1):1-12

Rallus crepitans (Gmelin): 2:1-20

Rangia cuneata (Sowerby, 1831): 2:63-73;
3(2):233-242; 6(2):189-197 (passim)

Rapana bezoar vaquerosensis: 2:85-85

Rapana imperialis: 2:85-85

Retusa: 5(2):185-196

Retusa canaliculata: 1:91

Retusa obtusa (Montagu): 5(2):197-214

Retusa truncata (Bruguiére, 1792):
5(2):243-258

Retusidae: 4(2):233; 5(2):243-258; S1:1-22

Retussa: S1:1-22

Rhinoclava (Proclava) kochii (Philippi,
1848): 2:1-20

Rhinocoela: 3(2):213-221 (passim)

Rhinoptera bonasus (Mitchell): S3:59-70

Rithropanopeus harrisii (Gould): S3:59-70

Rhizophora: 4(1):112

Rhizophora mangle Linné: 5(2):259-280

Rhizorus acuminatus Bruguiere:
5(2):185-196

Rhodope Koelliker, 1847: 6(1):57-68

(Rhombochiton) Berry, 1919: 3(1):63-82

Rhyssoplax Thiele, 1893: 6(1):115-130

Rhyssoplax affinis (Issel, 1869): 6(1):115-130

Rictaxis albus (Sowerby, 1873):
5(2):243-258

Rimula mexicana Berry, 1969: 3(1):63-82

Ringicula: $1:1-22

Ringicula buccinea (Brocchi): 5(2):185-196

Ringula turtoni Bartsch, 1915:
5(2):243-258

Ringiculidae: 5(2):243-258; S1:1-22

Risbecia pulchella (Riippell and Leuckart,
1828): 5(2):243-258

Rissoa albella Loven, 1846: 4(1):185-199
(passim)

Rissoa parva Da Costa: 5(2):303-306

Rissoacea H. and A. Adams, 1854:
3(2):223-231

Rissoella Gray, 1847: S1:1-22

Rissoella caribaea Rehder, 1943:
4(2):185-199

Rissoellidae Gray, 1850: S1:1-22

Rissoidae H. and A. Adams, 1854:
3(2):223-231; 4(2):235

Rissoina ambigua (Gould, 1849):
4(2):232-233

Rissoina bryerea (Montagu, 1803):
4(2):185-199

Rissoina catesbyana Orbigny, 1842:
4(2):185-199

Robastra gracilis (Bergh, 1877):
5(2):243-258

Robastra luteolineata (Baba, 1936):
5(2):243-258

Robsonella fontanianus (Orbigny):
6(2):207-211

Rossia Owen, 1835: 4(2):217-227

Rossia (Aust[rojrossia) australis Berry,
1918: 3(1):63-82

Rossia macrosoma (Delle Chiaje, 1829):
4(2):217-227

Rossia pacifica Berry, 1911: 2:91-92;
3(1):63-82

Rossia pacifica diegensis Berry, 1912:
3(1):63-82

Rostanga Bergh, 1879: 5(2):185-196

Rostanga muscula (Abraham, 1877):
5(2):243-258

Rostanga pulchra McFarland, 1905:
5(2):185-196, 197-214

Rostanga rubra (Risso, 1818): 5(2):185-196

Rostangidae Pruvot-Fol, 1954: 5(2):243-258

Rotella nana Lea, 1833: 4(1):39-42

Roxania Paetel, 1875: 5(2):185-196; S1:1-22

Roxania utriculus (Brocchi): 5(2):185-196

Roya \redale, 1912: 5(2):215-241

Roya spongotheras: 5(2):215-241

Rumina decollata (Linné, 1758): 1:23
(passim); 6(1):16

Runcina Forbes and Hanley, 1853:
5(2):185-196

Runcina coronata (Quatrefages, 1844):
5(2):185-196

Runcina ferruginea Kress: 5(2):185-196,
197-214

Runcina katipoides Miller and Rudman:
5(2):185-196

Runcina setoensis Baba: 5(2):197-214

Sacoglossa Ihering, 1876: 4(1):109-110;
5(2):243-258; $1:1-22

Saccostrea Dollfus and Dautzenberg,
1920: 4(2):157-162

Saccostrea cucullata (Born, 1778):
4(2):157-162

Saccostrea palmula (Carpenter, 1857):
4(2):157-162

Sagartia troglodytes (Price): 5(2):185-196

Salicornia: 2:1-20

Salinator: $1:1-22

Salmo salar (Linne): 5(2):125-128 (passim)

Salmo trutta Linne: 5(1):73-84; 5(2):125-128

Salvelinus fontinalis (Mitchell): 6(1):19-37

Salvia mellifera: 2:98

Samarangia quadrangularis Adams and
Reeve: S1:35-50

Sandalops ecthambus Berry, 1920:
3(1):63-82

Sandalops pathopsis Berry, 1920:
3(1):63-82

Sanguinolaria toulai Hertlein and Jordan,
1927: 4(1):1-12

Sargassum: 4(2):235; 5(2):259-280
(passim)

Saxidomus nuttalli: 4(2):241-242

Sayella: S1:1-22

Scaeurgus patagiatus Berry, 1913:
3(1):63-82; 6(2):207-211

Scaeurgus unicirrhus (Orbigny, 1840):
6:(2):207-211

Scalenostoma subulata (Broderip, 1832):
2:84

Scalptia mercadoi Old, 1968: 1:75-78

Scalptia nassa (Gmelin, 1791): 2:57-61

Scalptia scala (Gmelin, 1791): 2:57-61

Scalptia withrowi (Petit, 1976): 2:57-61

Scaphander: $1:1-22

Scaphander lignarius (Linne): 5(2):185-196

Scaphander punctostriatus (Mighels,
1841): 5(2):243-258

Scaphandridae: 4(2):233; 5(2):243-258;
$1:1-22

Scaphella contoyensis Emerson and Old,
1979: 1:75-78

Scaphopoda Bronn, 1862: 3(1):93-94

Scenedesmus: 4(1):81-88; S2:143-150

Scenella: 6(1):69-78

Schistosoma aematobium: 1:107

Schistosoma japonicum: 2:88

Schistosoma mansoni: 1:67-70, 106;
4(1):120; 5(1):85-90; S1:79-83

Schistosoma mansoni Puerto Rican PR-1:
1:106

Schistosoma mansoni Puerto Rican PR-2:
1:106

Schizochiton jousseaumei Smythe, 1982:
6(1):115-130

Schwartziella gracilis (Pease, 1861):
4(2):232-233

Scissurella lyra Berry, 1947: 3(1):63-82

Scissurella pseudoequatoria Kay, 1979:
4(2):232-233

Sclerodoris apiculata (Alder and Han-
cock, 1864): 5(2):243-258

Sclerodoris coriacea Eliot, 1904:
5(2):243-258

Scoloplos: 2:96

Scrobicularia: 3(2):213-221 (passim)

Scutopus: 6(1):57-68

Scutopus megaradulatus Salvini-Plawen:
6(1):57-68

Scyllaea pelagica Linné: 5(2):197-214

Scyllaeidae: 5(2):243-258

Searlesia dira (Reeve): 4(2):173-183
(passim)

Sebradoris crosslandi (Eliot): 5(2):197-214

Seguenzia (Jeffreys) Seguenza, 1876:
1:92

Seguenziacea: 1:92

Semele decisa: 4(2):241-242

Semibalanus balanoides: S1:111-116

Sepia: 4(2):217-227

Sepia chirotrema Berry, 1918: 3(1):63-82

250 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Sepia dannevigi Berry, 1918: 3(1):63-82

Sepia elegans Orbigny, 1835:
4(2):217-224

Sepia formosana Berry, 1912: 3(1):63-82

Sepia hedleyi Berry, 1918: 3(1):63-82

Sepia officinalis Linné, 1758: 2:91;
4(2):165-172, 217-227, 240, 241

Sepia orbignyana Ferussac, 1826: 2:91;
4(2):217-227

Sepiardium austrinum Berry, 1912:
3(1):63-82

Sepiardium nipponianum Berry, 1932:
3(1):63-82

Sepietta: 4(2):217-227

Sepietta oweniana: 2:90

Sepiodea Berry, 1920: 3(1):63-82

Sepiola: 4(2):217-227

Septemchiton Bergenhayn, 1955:
6(1):57-68

Septifer: 5(2):159-164

Serripes groenlandicus (Bruguierem,
1789): 2:94

Setoaeolis pilata (Gould): 5(2):287-292

Simpsonaias ambigua (Say, 1825):
2:85-86; 3(1):47-53; 6(1):19-37

Simpsoniconcha ambigua (Say, 1825):
3(1):105; 6(1):19-37

Simrothiella Pilsbry, 1898: S1:23-34

Simrothiellidae: $1:23-34

Sinonovacula: 5(2):159-164

Siphocyraea henekeni (Sowerby, 1850):
4(1):1-12

Siphonaria: 5(2):215-241; S1:1-22

Siphonaria alternata Say: S1:35-50

Siphonaria lessoni: 4(2):233

Siphonaria maura pica Sowerby, 1835:
4(1):1-12

Siphonaria williamsi Berry, 1969: 3(1):63-82

Siphonariidae: 2:88-89; S1:1-22

Skeletonema costatum (Greville): 4(1):81-88

Skenea (?) cyclostoma Berry, 1941:
3(1):63-82

Smaragdia viridis viridemaris Maury, 1917:
4(2):185-199

Smaragdinella: $1:1-22

Smaragdinella calyculata (Broderip and
Sowerby, 1829): 5(2):243-258

Smerinthus ocellatus: 5(2):185-196

Solariella carvalhoi: 3(1):101-102

Solatia Jousseaume, 1887: 2:57-61

Solatisonax |Iredale, 1931: 4(1):108-109

Solemya (Acharax) bartschi Dall, 1908:
$1:23-34

Solemya (Acharax) caribbaea Vokes:
$1:23-34

Solemya (Acharax) johnsoni Dall, 1891:
$1:23-34

Solemya agassizi Dall: S1:23-34

Solemya panamensis: S$1:23-34

Solemya reidi Bernard, 1980: 2:94

Solemya velum Say, 1822: S$1:23-34

Soleymidae H. and A. Adams, 1857
(1840): 4(1):111-112; S1:23-34

Solemyoidae Dall, 1889: 4(1):111-112

Solenogastres Gegenbaur, 1878: 4(1):107;
6(1):57-68

Solenosteira: 4(1):1-12

Solenosteira gatesi Berry, 1963: 3(1):63-82

Soletellina elongata Lamarck: S2:1-5
(passim)

Solivaga finschi (Thiele, 1910): 6(1):115-130

Sonorelix Berry, 1943: 3(1):63-82

Sonorella Pilsbry, 1900: 4(1):113-114

Sonorella anchana Berry, 1948: 3(1):63-82

Sonorella rooseveltiana Berry, 1917:
3(1):63-82

Sonorella strongiana Berry, 1948: 3(1):63-82

Sonorella virilis Pilsbry, 1905: 2:98

Spartina alternaflora Loiseleur-
Deslongchamps: 3(1):103

Sphaerium spp.: 2:86, 88; 5(1):21-30
(passim); S2:187-191, 193-201

Sphaerium corneum (Linné, 1758):
3(2):187-200, 201-212; 5(1):21-30
(passim), 41-48; S2:223-229

Sphaerium occidentale Prime, 1851:
3(2):187-200; S2:223-229

Sphaerium rhomboideum (Say, 1822):
5(1):31-39, 91-99, 105-124 (passim);
$2:223-229

Sphaerium rivicola: 3(2):187-200

Sphaerium scaldianum: 3(2):187-200

Sphaerium simile (Say, 1816): 5(1):31-39,
91-99, 105-124 (passim): S2:223-229

Sphaerium solidum: 3(2):187-200

Sphaerium striatinum (Lamarck, 1818):
3(2):187-200, 201-212 (passim);
4(1):116; 5(1):1-7, 31-39, 49-64,
105-124; S2:219-222, 223-229

Sphaerium suecicum: 3(2):187-200

Sphaerium transversum (Say, 1829):
5(1):41-48 (passim); S2:7-39

Sphincterochila aharonii (Kobelt): 6(1):16

Sphincterochila cariosa (Oliver): 6(1):16

Sphincterochila fimbriata (Bourguignat):
6(1):16

Sphincterochila prophetarum
(Bourguignat): 6(1):16

Sphincterochila zonata: 6(1):16

Spilogale putorius: 5(2):185-196

Spiraxidae: 1:97

Spiricella Rang, 1827: 5(2):215-241

Spirodon carinata Bruguiére: 3(2):169-177

Spirula Lamarck, 1799: 4(2):217-227

Spiruloidea Berry, 1920: 3(1):63-82

Spisula confraga (Conrad, 1833):
4(1):39-42

Spisula modicella (Conrad, 1833):
4(1):39-42

Spisula solidissima (Dillwyn, 1817): 1:13
(passim); 2:35-40; 3(2):135-142
(passim); S1:59-78

Spondylus nicobaricus Schreiber, 1793: 2:84

Spondylus ursipes Berry, 1959: 3(1):63-82

Spurilla neapolitana (delle Chiaje):
5(2):185-196

Squalus: 2:91-92

Spurwinkia salsa: 4(1):101-102

Stagnicola sp.: 1:97

Stagnicola elodes (Say, 1821): 5(1):9-19

Stagnicola palustris (Miller, 1776):
5(1):65-72 (passim)

Stauroteuthis (?) mawsoni Berry, 1917:
3(1):63-82

Stearnsium Berry, 1958: 3(1):63-82

Stenomena: S2:69-81

Stenoplax (Maugerella) conspicua
sonorana Berry, 1956: 3(1):63-82

Stenoplax (Stenoradisia) heathiana Berry,
1946: 3(1):63-82

Stenoplax circumsenta Berry, 1956:
3(1):63-82

Stenoplax histrio Berry, 1945: 3(1):63-82

Stenoplax isoglypta Berry, 1956:
3(1):63-82

Stenotrema fraternum (Say, 1824): 1:98

Stephanodiscus: S2:167-178

Stephanoteuthis Berry, 1909: 3(1):63-82

Stephanoteuthis hawaiiensis Berry, 1909:
3(1):63-82

Stichodactyla helianthus (Ellis, 1768):
1:1-12

Stichopus chloronatus: 2:83

Stiliger: S1:1-22

Stiliger fuscovittatus Lance: 5(2):197-214

Stiliger ornatus Ehrenberg, 1831:
5(2):243-258

Stiligeridae: 5(2):243-258, 259-280;
$1:1-22

Stoloteuthinae Berry, 1914: 3(1):63-82

Stoloteuthis iris Berry, 1909: 3(1):63-82

Stoloteuthis nipponensis Berry, 1911:
3(1):63-82

Striostrea Vialov, 1936: 4(2):157-162

Striostrea circumpicta (Pilsbry, 1904):
4(2):157-162

Striostrea margaritacea (Lamarck, 1819):
4(2):157-162

Striostrea prismatica (Gray, 1825):
4(2):157-162

Striostrea (Parastriostrea): 4(2):157-162

Striostrea (Parastriostrea) mytiloides
(Lamarck, 1819): 4(2):157-162

Striostreini: 4(2):157-162

Strombidae Rafinesque, 1815: 4(1):109-110

Strombina Morch, 1852: 4(1):1-12

Stromboli Berry, 1954: 3(1):63-82

Strombus Linné, 1758: 4(2):157-162
(passim); 185-199 (passim)

Strombus gigas Linné, 1758: 3(2):223-231

Strombus lineolatus Gray, 1828: 2:1-20
(passim)

Strombus (Tricornis) costatus (Gmelin,
1791): 4(1):108

Strombus (Tricornis) leidyi (Heilprin, 1887):
4(1):108

Strombus (Tricornis) mayacensis (Tucker
and Wilson): 4(1):108

Strombus oldi Emerson, 1965: 1:75-78

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 251

Strophitus rugosus Dall, 1905: 1:43-50

Strophitus rugosus (Swainson, 1822):
6(1):19-37

Strophitus subvexus (Conrad, 1834):
4(1):21-23

Strophitus undulatus (Say, 1817): 1:28,
43-50; 3(1):41-45; 4(1):41-45;
6(1):19-37

Strophitus undulatus tennessensis (Lea,
1840): 4(1):117-118

Strophitus undulatus undulatus (Say,
1817): 1:51-60; 2:85-86; 3(1):47-53,
105; 5(2):165-171

Struthiolariidae: $1:35-50

Stylocheilus longicauda (Quoy and
Gaimard, 1824): 5(2):243-258

Stylochus: S3:59-70

Stylochus ellipticus (Gould): S3:59-70

Stylpopodium zonale (Lamouroux) Papen-
fuss: 5(2):259-280 (passim)

Succinea ovalis: 1:97-98

Susania Gray, 1857: 5(2):215-241

Syllis: 2:29

Symplectoteuthis oualaniensis (Lesson,
1830): 2:51-56

Synedra: S2:167-178

Syrnolopsidae: 3(2):223-231

Systellommatophor: $1:1-22

Takydromus tachydromoides oldi Walley,
1958: 1:75-78

Tambja capensis Bergh, 1907:
5(2):243-258

Tambja morosa (Bergh, 1877):
5(2):243-258

Tarebia granifera (Lamarck, 1819): 1:95-96

Tautogolabris adspersus (Walbaum):
5(2):287-292

Tegula sp.: 1:102; 2:41-50; 4(1):1-12

Tegula pfeifferi: 4(2):165-172

Teinostoma nana (Lea, 1833): 4(1):39-42

Teleoteuthis compacta Berry, 1913:
3(1):63-82

Telescopium Montfort, 1910: 2:1-20

Telesto: 5(2):185-196

Telledorella Berry, 1963: 3(1):63-82

Telledorella cristulata Berry, 1963:
3(1):63-82

Tellina Linné, 1758: 3(2):213-221 (passim)

Tenellia adspersa (Nordmann):
5(2):287-292

Tenellia pallida (Nordmann): 4(2):205-216
(passim); 5(2):197-214

Terebra burckhardti Hertlein and Jordan,
1927: 4(1):1-12

Terebra (Strioerebrum) danai Berry, 1958:
3(1):63-82

Terebra (Strioerebrum) fitchi Berry, 1958:
3(1):63-82

Terebra (Strioerebrum) puncturosa Berry,
1959: 3(1):63-82

Terebralia Swainson, 1840: 2:1-20

Terebralia palustris: 2:1-20

Teredinidae: 3(1):85-88

Teredo bartschi Clapp: 4(1):89-99;
$1:101-109; S2:203-209

Teredo furcifera von Martens: S1:101-109

Teredo navalis Linné: 3(1):85-88;
4(1):89-99; S1:101-109

Tergipedidae: 5(2):243-258

Tergipes tergipes Forskal, 1779:
5(2):185-196, 197-214, 243-258

Teskeyostrea: 4(2):157-162

Teskeyostrea weberi Olsson, 1951:
4(2):157-162

Testacea: 2:82

Tethyidae: 5(2):243-258

Tethys fimbria Linné: 5(2):197-214

(Teuthidiscus) Berry, 1918: 3(1):63-82

Teuthowenia (Ascoteuthis) corona Berry,
1920: 3(1):63-82

Thaididae: 3(1):11-26; 4(1):109-110

Thais Roding, 1798: 3(2):213-221
(passim); 4(1):110; 5(2):293-301
(passim)

Thais emarginata (Deshayes, 1839): 1:105

Thais deltoidea (Lamarck, 1822): 1:8

Thais floridana haysae Clench, 1927:
6(2):189-197

Thais haemastoma (Linné, 1758): 2:63-73;
6(1):17; S1:35-50; 6(2):189-197

Thais haemastoma canaliculata (Gray,
1839): 2:63-73; 6(2):189-197

Thais haemastoma floridana (Conrad,
1837): 6(2):189-197

Thais haysae Clench, 1927: 6(2):189-197

Thais lamellosa (Gmelin): 6(1):178

Thais lapillus (Linné, 1758): 2:63-73;
4(2):165-172

Thais nodosa: 4(1):110

Thais nodosa mevetricula: 3(1):101-102

Thais savignyi: 4(1):109-110

Thalamita crenata: 4(1):112

Thalassia testudinum (KOnig, 1805):
4(2):185-199; 5(2):259-280

Thalassoma bifasciatum (Bloch, 1791): 1:8

Theba pisana (Miller, 1774): 1:104,
104-105; 6(1):16

Thecacera pacifica (Bergh, 1884):
5(2):243-258

Thecacera pennifera (Montagu, 1804):
5(2):197-214

Thecacera pennigera (Montagu, 1804):
5(2):243-258

Thecosomata: $1:1-22

Theodoxia fluviatilis (Linné, 1758):
5(1):65-72 (passim)

Theodoxus: 4(1):1-12

Theodoxus fluviatilis (Linné, 1758):
4(1):185-199 (passim)

Thiaridae: 3(2):223-231

Thordisa filix Pruvot-Fol: 5(2):197-214

Thorunna clitonata (Bergh): 5(2):197-214

Thorunna decussata (Risbec):
5(2):197-214

Thorunna norba (Marcus and Marcus):
5(2):197-214

Thracia phaseolina Lamarck: S1:35-50

Thracia pubescens: 2:35-40

Thraciaciidae: 2:35-40

Thraciidae Stoliczka, 1870: S1:35-50

Thunnus alalunga: 4(2):241

Thyca (Bessomia) callista Berry, 1959:
3(1):63-82

Thyrasira: $1:23-34

Thyrasiridae: 2:29; $1:23-34

Tiariturris Berry, 1958: 3(1):63-82

Tiariturris spectabilis Berry, 1958:
3(1):63-82

Tiphyocerma Berry, 1958: 3(1):63-82

Tiphyocerma preposterum Berry, 1958:
3(1):63-82

Tivela scarificata Berry, 1940: 3(1):63-82

Toledonia: $1:1-22

Tonicella insignis Reeve, 1847:
6(1):141-151

Tonicella marmorea (Fabricius, 1780):
6(1):69-78, 153-159

Tonicella rubra (Linné, 1767): 6(1):69-78

Tonicia Gray, 1847: 6(1):115-130

Tonicia ptygmata Rochebrune, 1883:
6(1):115-130

Tonicia (Lucilina) carnosa Kaas, 1979:
6(1):115-130

Tonicia (Lucilina) sueziensis (Reeve,
1847): 6(1):115-130

Toniciinae Pilsbry, 1893: 6(1):115-130

Tornatina decurrens Verrill and Bush,
1900: 3(1):93

Tornatina inconspicusa Olsson and
McGinty, 1958: 3(1):93

Tornatina liratispira E. A. Smith, 1872:
3(1):93

Toxolasma cylindrella (Lea, 1868): 6(1):19-37

Toxolasma cylindrellus (Lea, 1868):
6(2):165-178

Toxolasma livida Rafinesque, 1831:
6(1):19-37

Toxolasma lividium (Rafinesque, 1831):
6(2):165-178

Toxolasma lividus (Rafinesque, 1831):
3(1):41-45, 104; 6(2):165-178

Toxolasma lividus glans (Lea, 1831):
6(1):19-37

Toxolasma lividus lividus (Rafinesque,
1831): 6(1):19-37

Toxolasma parva: 6(1):19-37

Toxolasma parvus (Barnes, 1823): 1:51-60;
2:86

Toxolasma pullus (Conrad, 1838): 1:61-68

Toxolasma texasensis (Lea, 1857):
4(1):21-23; 6(1):19-37

Trachycardium Morch, 1853: 4(1):1-12

Transennella caryonautes Berry, 1963:
3(1):63-82

Transennella tantilla (Gould, 1852): 2:94

Trapania: 5(2):243-258

Trapania maculata Haefelfinger:
5(2):185-196, 197-214

Tremoctopus: 4(2):217-227

202 AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988

Trialatella Berry, 1964: 3(1):63-82

Trialatella cunninghamae Berry, 1964:
3(1):63-82

Trichoptera: S2:69-81

Tricolia affinis affinis (C. B. Adams, 1850):
4(2):185-199

Tricolia affinis cruenta Robertson, 1958:
4(1):185-199 (passim)

Tricolia bella (M. Smith, 1937): 4(2):185-199

Tricolia thalassicola Robertson, 1958:
4(2):185-199

Tricolia variabilis (Pease, 1861):
4(2):232-233

Tricula Benson, 1843: 2:88

Triculinae Annandale, 1924: 3(1):96

Tridachia crispata Morch, 1863: 4(2):232;
5(2):197-214

Tridacna sp.: 2:83

Tridacna maxima (Roding, 1798): 1:18
(passim)

Trigona pellucida Perry, 1811: 2:57-61

Trigonaphora withrowi Petit, 1976: 2:57-61

Trigonioida: 4(1):13-19

Trigonostoma Blainville, 1827: 2:57-61

Trigonostoma antiquata (Hinds, 1843):
2:57-61

Trigonostoma lamellosa (Hinds, 1843):
2:57-61

Trigonostoma pellucida (Perry, 1811):
2:57-61

Trigonostoma scalare (Gmelin, 1791):
2:57-61

Triodopsis Rafinesque, 1819: 2:97-98

Triodopsis albolabris (Say, 1816): 1:98;
2:98; 6(1):16

Triodopsis albolabris alleni (‘Wetherby’
Sampson, 1881): 1:97-98

Triodopsis fosteri: 2:98

Triodopsis multilineata (Say, 1821): 1:97-98

Triodopsis tridentata tridentata (Say, 1816):
1:98

Triopha catalinae (Cooper): 5(2):197-214,
5(2):287-292

Triopha carpenteri Stearns: 5(2):185-196

Triopohridae: $1:1-22

(Tripoplax) Berry, 1919: 3(1):63-82

Trippa spongiosa (Kelaart): 5(2):197-214

Tritogonia Agassiz, 1852: 4(1):117-118

Tritogonia verrucosa (Rafinesque, 1820):
1:29, 43-50, 51-60, 71-74; 2:85-86;
3(1):47-53; 4(1):21-23; 5(2):165-171;
6(1):19-37

Tritonia: 2:78; 5(2):243-258

Tritonia diomeda Bergh: 1:13 (passim);
2:78; 5(2):185-196, 197-214

Tritonia festiva (Stearns): 5(2):197-214

Tritonia hombergi Cuvier: 4(1):103-104;
4(2):205-216, 235; 5(2):197-214

Tritonia nilsodhneri Marcus, 1983:
5(2):185-196, 243-258

Tritoniidae: 5(2):243-258

Tritoniopsis cincta Pruvot-Fol: 5(2):197-214

Tritonium viridulum Fabricius, 1780: 2:57-61

Trivia exigua Gray, 1930: 4(2):232-233

Trochacea: 3(1):104; $1:23-34

Trochidae: 3(1):95; 4(1):109-110; S1:1-22

Trochita radians ‘Lamarck’ Arnold and
Anderson, 1907: 4(1):1-12

Trochita spirata (Forbes, 1872): 4(1):1-12

Trochita trochiformis (Born, 1778): 4(1):1-12

Trochostylifer sp.: 2:83

Trochus erythraeus: 4(1):109-110

Trophon acanthodes Watson, 1882:
3(1):101-102

Trophon aculeatus Watson, 1882: 3(1):11-26

Trophon bahamondei McLean and
Andrade, 1982: 3(1):11-26

Trophon longstaffi Smith, 1904: 3(1):11-26

Trophon (Pagodula) acanthodes (Watson,
1882): 3(1):101-102

Trophon shackeltoni Hedley, 1911: 3(1):11-26

Trophon truncatus Strom, 1768: 3(1):11-26

Trophoninae: 3(1):11-26

Truncilla donaciformis (Lea, 1828):
1:43-50, 51-60, 71-74; 3(1):105;
6(1):19-37

Truncilla truncata Rafinesque, 1820: 1:29,
43-50, 51-60, 71-74; 2:85-86; 3(1):105;
6(1):19-37

Truncilla vermiculata (Rafinesque, 1820):
6(1):19-37

Tubastraea coccinea Lesson: 5(2):197-214

Tubularia spp. 5(2):185-196

Turbinaria: 5(2):185-196

Turbinella angulata (Lightfoot, 1786):
4(1):113

Turbinidae: 4(1):109-110

Turbo radiatus: 4(1):109-110

Turbonilla Risso, 1826: $1:1-22

Turbonilla vineae Bartsch, 1909: S1:1-22

Turcica admirabilis Berry, 1969: 3(1):63-82

Turridae Swainson, 1840: 3(1):98;
$1:23-34

Turrigemma Berry, 1958: 3(1):63-82

Turrigemma torquifer Berry, 1958:
3(1):63-82

Turritella sp.: 2:84-85

Turritella abrupta Spieker: 2:84-85;
4(1):1-12

Turritella altilira Conrad, 1857: 4(1):1-12

Turritella anactor Berry, 1957: 3(1):63-82

Turritella bifastigata Nelson: 4(1):1-12

Turritella bosei Hertlein and Jordan, 1927:
4(1):1-12

Turritella communis: 3(2):179-186 (passim)

Turritella costaricensis Olsson, 1922:
4(1):1-12

Turritella crocus Cooke, 1919: 4(1):1-12

Turritella inezana Conrad: 4(1):1-12

Turritella inexana bicarina Loel and Corey,
1932: 4(1):1-12

Turritella ocoyana Conrad: 4(1):1-12

Turritella orthosymmetra Berry, 1953:
3(1):63-82

Turritellidae Clarke, 1851: 3(1):95; S1:1-22,
35-50

Turveria Berry, 1956: 3(1):63-82

Turveria ecopendema Berry, 1956:
3(1):63-82

Tylodina Rafinesque, 1819: 5(2):215-241

Tylodina alfredensis Turton, 1932:
5(2):243-258

Tylodina citrina Joannis, 1834:
5(2):215-241

Tylodina corticalis (Tate): 5(2):215-241

Tylodina duebeni Loven, 1846: 5(2):215-241

Tylodina fungina: 5(2):215-241

Tylodina perversa (Gmelin): 5(2):215-241

Tylodinella Mazzarelli, 1898: 5(2):215-241

Tylodinella trinchesii Mazzarelli, 1898:
5(2):215-241

Tylodinidae Gray, 1847: 5(2):215-241

Tympanotonus fascatus (Linné, 1758): 2:1-20

Typhina riosi: 3(1):101-102

Udotea conglutinata (Ellis and Solander)
Lamouroux: 5(2):259-280

Ulva: 5(2):287-292

Ulva lactuca: 1:92

Umbonium Link: 3(1):95; 4(1):109

Umba limi (Kirtland): 5(1):73-84

Umbraculacea Dall, 1889: 5(2):215-241

Umbraculidae Dall, 1889: 5(2):215-241,
243-258; S1:1-22

Umbraculum Schumacher, 1817:
5(2):215-241, 243-258; $1:1-22

Umbraculum sinicum (Gmelin, 1783):
5(2):243-258

Umbraculum umbraculum (Lightfoot):
5(2):215-241

Umbrella Lamarck, 1819: 5(2):215-241

Undulostrea: 4(2):157-162

Undulostrea megodon (Hanley, 1846):
4(2):157-162

Unela glandulifera (Kowalevsky):
5(2):303-306

Unela nahantensis Doe: 3(1):27-31
(passim); S1:35-50

Unio Philipsson, 1788: 4(2):157-162

Unio moestus Lea: 6(2):165-178

Unio pictorum Linné, 1758: 3(2):233-242

Unio (Toxolasma) cylindrellus Lea, 1868:
6(2):165-178

Uniomerus declivus (Say, 1831):
4(1):21-23; 6(1):19-37

Uniomerus tetralasmus (Say, 1830):
4(1):21-23; 6(1):19-37

Uniomerus tetralasmus manubius (Gould,
1855): 2:86

Union douglasiae (Gray, 1833): 5(1):91-99

Unionacea Fleming, 1828: 3(2):201-212;
4(1):13-19

Uniondae Fleming, 1828: 6(2):179-188

Unionidae, Unspecified: 1:93, 93-94, 97;
2:86, 86-87; 3(1):106, 106-107;
4(1):61-79, 101; 4(2):157-162 (passim);
$2:1-5

Upogeba: 1:90-91

Urosalpinx cinerea (Say, 1822): 2:63-73;
4(2):165-172; S1:111-116; S3:59-70

AMER. MALAC. BULL. TAXONOMIC INDEX: 1983 - 1988 253

Urosalpinx cinerea follyensis Baker, 1951:
2:63-73

Urosalpinx perrugata (Conrad, 1846):
4(1):185-199 (passim)

Utriculastra Thiele, 1925: 4(1):39-42

Vallisneria americana: 5(1):73-84

Valvata Miller, 1774: S1:1-22

Valvata piscinalis (Muller, 1774):
3(2):243-265

Valvata tricarinata (Say, 1817): 5(1):9-19,
31-39, 105-124 (passim)

Valvatacea: 3(2):223-231; S1:1-22

Valvatidae Gray: 3(2):223-231; $1:1-22

Vampyroteuthis Chun, 1903: 4(2):217-227

Vampyroteuthis infernalis Chun, 1903:
4(2):217-227

Vanikoro cancellata (Lamarck, 1822):
4(2):232-233

Vaucheria: 5(2):197-214, 259-280

Vasinae H. and A. Adams, 1854: 3(1):11-26

Vasum pufferi: 2:84-85

Vayssieridae: 5(2):243-258

Velella: 5(2):185-196

Velesunio: 4(1):13-19

Vema: 6(1):69-78

Venustaconcha ellipsiformis ellipsiformis
(Conrad, 1836): 5(2):165-171

Vermes: 2:82

Vermetidae: 3(1):95

Vermetus contortus (Carpenter, 1857):
4(1):1-12

Veronicellidae: S1:1-22

Verrilliteuthis Berry, 1916: 3(1):63-82

Verticordiidae Stoliczka, 1971 (sic): S1:35-50

Verticumbo Berry, 1940: 3(1):63-82

Verticumbo charybdis Berry, 1940:
3(1):63-82

Vertigo allyniana Berry, 1919: 3(1):63-82

Vertigo allyniana xenos Berry, 1919:
3(1):63-82

Vertigo modesta micorphasma Berry,
1919: 3(1):63-82

Vesicomya Dall, 1886: S1:23-34

Vesicomya caudata Boss: $1:23-34

Vesicomya cordata Boss: $1:23-34

Vesicomyidae Dall and Simpson, 1901:
1:101; 3(1):95-96; S1:23-34

Vestimentifera: $1:23-34

Vexillum (Pusia) chickcharneorum Lyons
and Kaicher, 1978: 4(1):113

Vibrio alginolyticus: 2:93-94

Vibrio damsela: 2:93-94

Vibrio parahaemolyticus: 2:93-94

Villosa Frierson, 1927: 4(1):117-118;
6(2):165-178

Villosa fabalis (Lea, 1831): 1:43-50;
3(1):105; 6(1):19-37

Villosa iris (Lea, 1830): 1:43-50; 3(1):41-45,
105; 6(1):19-37; 6(2):165-178

Villosa iris iris (Lea, 1830): 2:85, 85-86;
3(1):47-53; 5(2):165-171

Villosa lienosa (Conrad, 1834): 1:29;
3(1):47-53; 4(1):21-23; 6(1):19-37

Villosa nebulosa (Conrad, 1834): 1:43-50;
3(1):104; 5(1):1-7; 6(1):19-37

Villosa ogeecheensis (Conrad, 1834):
1:61-68

Villosa ortmanni (Walker, 1925): 1:29

Villosa perpurpurea (Lea, 1861): 6(2):165-178

Villosa picta (Lea, 1834): 6(1):19-37

Villosa taeniata (Conrad, 1834): 1:43-50;
4(1):25-37; 6(1):19-37

Villosa taeniata picta (Lea, 1834):
6(1):19-37

Villosa taeniata punctata (Lea, 1865):
6(1):19-37

Villosa taeniata taeniata (Conrad, 1834):
6(1):19-37

Villosa teneltus (Rafinesque, 1831):
6(1):19-37

Villosa trabalis (Conrad, 1834): 1:27-30;
4(1):25-37; 6(1):19-37; 6(2):165-178

Villosa trabalis perpurpurea (Lea, 1861):
6(1):19-37

Villosa vanuxemensis (Lea, 1838):
3(1):41-45; 6(1):19-37; 6(2):165-178

Villosa vanuxemi (Lea, 1838): 1:43-50;
3(1):104; 4(1):25-37; 5(1):1-7;
6(2):179-188

Villosa vibex (Conrad, 1834): 6(1):19-37

Villosa villosa (Wright, 1898): 1:95;
4(1):117; 4(2):231

Virgularia: 5(2):197-214

Viriola abbotti (Baker and Spicer, 1935):
2:84

Vitrea orotis Berry, 1930: 3(1):63-82

Vitreolina sp.: 2:83

Viviparacea Gray, 1847: 3(2):223-231

Viviparidae Gray, 1847: 3(1):107;
3(2):223-231

NEW TAXA DESCRIBED IN THE

AMERICAN MALACOLOGICAL BULLETIN

Acanthochitona ferreirai Lyons, 1988: 6(1):85-86, Figs. 19-24 (Punta Mala, Panama).
Acanthochitona lineata Lyons, 1988: 6(1):90-92, Figs. 42-51 (Silver Cove Canal, Freeport, Grand Bahama Island).
Acanthochitona roseojugum Lyons, 1988: 6(1):98-100. Figs. 82-92 (Bartlett Hill, Eight Mile Rock, Grand Bahama Island).

Acanthochitona venezuelana Lyons, 1988: 6(1):96-98, Figs. 73-80 (Isla de Margarita, Venezuela).
Acanthochitona woodwardi Kaas and Van Belle, 1988, 6(1):126-127, Figs. 51-60 (Qatar, Dasa).

Viviparus Montfort, 1810: 3(2):269-272

Viviparus bengalensis: 3(2):223-231

Viviparus contectoides (Binney): 6(1):17

Viviparus georgianus (Lea): 3(2):268;
5(1):9-19

Viviparus melleatus (Reeve, 1863):
3(2):223-231

Vivaparus viviparous: 3(2):179-186
(passim), 223-231

Volsella sacculifer Berry, 1953: 3(1):63-82

Voluta cancellata Linné, 1767: 2:57-61

Voluta nassa Gmelin, 1791: 2:57-61

Voluta reticulata Linné, 1767: 2:57-61

Voluta scabriculus (Linné, 1758): 2:57-61

Volutidae: 3(1):101-102

Volvatella: $1:1-22

Volvatella bermudae Clark: 5(2):259-280

Volvatella laguncula Sowerby, 1894:
5(2):243-258

Volvatellidae: S1:1-22

Volvulidae: 4(2):233

Vorticella sp.: 3(2):151-168

Westraltrachia |redale, 1933: 1:98-99

Williamia Monterosato, 1844: 2:88-89;
5(2):215-241; $1:1-22

Williamia gussonii (Costa, 1829): 2:88-89

Woodbridgea Berry, 1953: 3(1):63-82

Woodbridgea williamsi Berry, 1953:
3(1):63-82

Xenia: 5(2):185-196

Xerarionta: 3(1):102-103

Xerocrassa seetzeni (Pfeiffer): 6(1):16

Ximeniconus Emerson and Old, 1962:
1:75-78

(Xiphpiozona), Lepidopleurus Berry, 1919:
3(1):63-82

Yoldia hyperborea ‘Loven’ Torell, 1859:
2:94

Zaccatrophon Hertlein and Strong, 1951:
3(1):11-26

Zebina browniana (Orbigny, 1842):
4(2):185-199

Zemelanopsis: 2:1-20

Zidoninae: 3(1):11-26

Zostera marina Linne: 5(2):185-196

Zyzzyzus spongicola (von Lendenfeld):
5(2):185-196

Acanthochitona worsfoldi Lyons, 1988: 6(1):92-94, Figs. 57-65 (Silver Cove Canal, Freeport, Grand Bahama Islanq).
Acanthochitona zebra Lyons, 1988: 6(1):105-107, Figs. 115-127 (Silver Cove Canal, Freeport, Grand Bahama Island).

Anidolyta Willan, 1988: 5(2):232-233, Tylodina duebeni Lovén, 1846, type species by designation.

Notoplax (Notoplax) arabica Kaas and Van Belle, 1988, 6(1):127-128, Figs. 61-72 (Kuwait Bay, Kuwait, on rocks and dead shells, intertidal).

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Abaco, Bahama Islands

Acanthochitona andersoni, A.
pygmaea: 6(1):79-114

Aden

Chiton (Chiton) peregrinus,
Ischnochiton (Ischnochiton) yerbury!:
6(1):115-130

Africa

Acanthopleura brevispinosa:
6(1):115-130. Acteon fortis:
5(2):243-258. Aeolidiella indica:
2:95-96. African Great Lakes:
5(1):85-90. Algeria: 2:88-89; 5(1):85-90.
Amblychilepas: 2:21-34. Ancylus
drouetianus, A. gussonii: 2:88-89.
Aplysia dactylomela, A. juliana:
5(2):243-258. Bellamya, B. capillata,
B. jeffreysi, B. unicolor: 4(1):107.
Berthella plumula: 5(2):197-214.
Biomphalaria alexandrina: 1:107.

B. choanomphala: 5(1):85-90.

B. glabrata: 1:106-107. B. pfeifferi,

B. smithii, B. stanleyi: 5(1):85-90.

B. straminea: 1:106-107. B. sudanica:
5(1):85-90. Brondelia drouetiana,

B. gussonii 2:88-89. Bulinus natalensis,

B. tropicus: 1:96, 106-107. B. truncatus:

1:106-107; 5(1):85-90. Buccinum
piscatorium: 2:57-61. Caelatura:
4(1):107. Cancellaria (Bivetiella)
cancellata, C. lamellosa, C. (Solatia)
piscatoria: 2:57-61. Cape of Good
Hope: 6(1):115-130. Ceratophyllidia
africana, Chromodoris hamiltoni, C.
vicina: 5(2):243-258. Corbicula
aegyptica, C. africana, C. agrensis,
C. artini, C. astartina, C. australis, C.
cunningtoni, C. fischeri, C. fluminalis,
C. fluminea, C. kirkii, C. lamarckiana,
C. oliphantensis, C. pusilla, C. radiata,
C. sikarae, C. subradiata, C.
tanganyicensis: S2:113-124. Cuthona
kanga, Dolabrifera dolabrifera, East
Africa, Favorinus ghanensis:
5(2):243-258. Fissurella haintula:
2:21-34. Ghana: 5(2):185-196, 243-258.
Glossodoris, Godiva quadricolor:
5(2):243-258. Hydrobia aponensis:
5(1):85-90. Hypselodoris tema:
5(2):185-196. Jorunna zania:
5(2):243-258. Kenya: 6(1):115-130.
Lake Albert, Lake Edward:
5(1):85-90. Lake Malawi: 4(1):107.
Lake Victoria: 4(1):107; 5(1):85-90.
Lake Tanganyika: 4(1):107. Mazoe
Dam: 1:106-107. Melanoides tuber
culata, Melanopsis, Mercuria confusa,
M. punica: 5(1):85-90. Mohari For
mation: 4(1):107. Mozambique: 2:57-61;
6(1):115-130. Murex scala: 2:57-61.

GEOGRAPHIC INDEX

Natal: 6(1):115-130. Neothauma
tanganyicense: 4(1):107. Onithochiton
literatus, O. wahlbergi: 6(1):115-130.
Opisthobranchia: 2:95-96. Paleontology:
4(1):107. Panacca. P. africana, P
locardi: 3(1):103-104. Perna perna:
5(2):159-164. Pliodon, P ovata, P
spekii: 4(1):107. Pleurobranchus
brockii, P. tarda, Prutfolia pselliotes:
5(2):243-258. Pupillaea aperta:
2:21-34. Scalptia scala: 2:57-61.
Sclerodoris coriacea: 5(2):243-258.
Schistosoma mansoni: 5(1):85-90.
Siphonariidae: 2:88-89. South Africa:
5(2):197-214; 6(1):115-130. Tanzania,
Thecacera pennigera: 5(2):243-258.
Trigonaphora withrowi: 2:57-61.
Tunisia: 5(1):85-90. Viviparidae:
4(1):107. Voluta cancellata: 2:57-61.
West Africa: 5(2):243-258. Williamia
gussonil: 2:88-89. Zimbabwe:
1:106-107; 5(1):85-90

African Great Lakes

Biomphalaria choanomphala, B.
smithii, B. stanleyi, B. sudanica,
Schistosoma mansoni: 5(1):85-90

Agua Fria River, AZ

Corbicula fluminea: S2:7-39

Aille River, Republic of lreland

Ancylus fluviatilis: 5(1):105-124

Al Bastan Island, Oman

Callistochiton adenensis, Chiton
(Chiton) peregrinus: 6(1):115-130

Alabama (AL)

Actinonaias carinata, A. pectorosa,
Alasmidonta calceolus, A. marginata,
A. minor, Amblema costata, A.
plicata, Anculosa praerosa, Anodonta
grandis: 1:43-50. Big Cedar Creek,
Big Nance Creek, Black Warrior
River, Buck Creek, Burnt Corn Creek,
Cahaba River: S2:7-39. Carunculina
lividus, C. moesta, C. moesta cylin-
drella: 1:43-50. Cedar Creek, Chatta-
hoochee River, Choctawahatchee
River, Conecuh River: S2:7-39. Con-
rdailla caelata: 1:43-50. Coosa River,
Corbicula fluminea: S2:7-39. C.
manilensis: 1:43-50. Cypress Creek,
Dauphin Island, Drivers Branch:
$2:7-39. Dromus dromas, Dysnomia
biemarginata, D. brevidens, D. cap-
saeformis, D. florentina, D. haysiana,
D. torulosa, D. triquetra: 1:43-50. Elk
River: 1:43-50. Elk River: 1:43-50;
S2:7-39. Elliptio crassidens, E.
dilatatus: 1:43-50. Escambia River,
Flint River: S2:7-39. Fusconaia
barnesiana, F. barnesiana bigbyensis,
F. cuneolus, F. edgariana, F.

subrotunda: 1:43-50. Gantt Lake:
$2:7-39. Goniobasis laquetra:
1:43-50. Graptemys pulchra, Indian
Creek: S2:7-39. lo verrucosa lima:
1:43-50. Lampsilis altilis: 1:94. L.
anodontoides, L. fasciola, L. ovata,
L. ovata ventricosa: 1:43-50. L.
perovalis: 1:94. Lasmigona com-
planata, L. costata, Lastena lata,
Leptodea fragilis, Lexingtonia
dolabelloides, L. dolabelloides con-
radi: 1:43-50. Limestone Creek:
$2:7-39. Lithasia verrucosa lima:
1:43-50. Little Cypress Creek, Little
Uchee Creek, Locust Fork: S2:7-39.

Margaritifera margaritifera: 4(1):13-19.
Medionidus conradicus, Megalonaias

gigantea: 1:43-50. Mobile River
System: 1:94; S2:7-39. Mud Creek,
Murder Creek, Neely Henry Lake,
North River: S2:7-39. Obliquaria
reflexa, Obovaria subrotunda,

O. subrotunda lens: 1:43-50.
Okatuppa Creek, Paint Rock River,
Pea River, Peckerwood Creek:
S2:7-39. Pegias fabula: 1:43-50.
Piney Creek: S2:7-39. Plagiola
lineolata, Pleurobema cordatum, P
oviforme, P oviforme argentium,
Pleurocera canaliculatum,
Ptychobranchus fasciolaris, P.
subtentum, Quadrula cylindrica, Q.
intermedia, Q. metanevra, Q.
pustulosa, Q. quadrula: 1:43-50.
Santa Bogue Creek, Saugahatchee
Creek, Second Creek, Sepulga
River: S2:7-39. Strophitus rugosus,
S. undulatus: 1:43-50. Sucarnochee
Creek, Tallapoosa River, Terrapin
Creek, Tombigbee River, Town

Creek: S2:7-39. Tritogonia verrucosa.

Truncilla donaciformis, T. truncata:

1:43-50. Tubbs Creek, Uchee Creek:

S2:7-39. Villosa fabalis, V. iris, V.
nebulosa, V. taeniata, V. vanuxemi:
1:43-50.
Alamo Canal, CA
Corbicula fluminea: S2:7-39
Alaska (AK)
Asterias amurensis: 2:94. Bering
Sea: 1:105. Macoma calcarea,
Mya truncata, Norton Sound: 2:94.
Nucella emarginata: 1:105. Serripes
groenlandicus, Yoldia hyperborea:
2:94. Thais emarginata:
1:105
Alberta, Canada
Cionella lubrica: 3(1):27-32
Aleutian Trench
Prochaetodermatidae: 3(1):97

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 205

Algeria
Ancylus drouetianus, A. gussonii,
Brondelia drouetiana, B. gussonii:
2:88-89. Bulimus truncatus:
5(1):85-90. Siphonariidae, Williamia
gussonii: 2:88-89.
All American Canal, CA
Corbicula fluminea: S2:7-39
Allan Branch, MS
Corbicula fluminea: S2:7-39
Altamaha River, GA
Corbicula: S2:1-5. C. fluminea:
S2:7-39. Elliptio shepardiana: 3(1):94
Amite River, MS
Corbicula fluminea: S2:7-39
Anaheim Bay, Los Angeles, CA
Cerithidea californica: 2:1-20
Anclote Key, FL
Acanthochitona pygmaea: 6(1):79-114
Andaman Islands
Ischnochiton (Ischnochiton) winck-
worthi: 6(1):115-130
Andros Island, Bahama Islands
Acanthochitona roseojugum,
Choneplax lata: 6(1):79-114
Angelina River, TX
Corbicula fluminea: S2:7-39
Angostura Formation, Ecuador
Turritella inezana, T. ocoyana:
4(1):1-12
Anguilla
Paziella pazi 3(1):11-26
Antarctica
Lissarca notorcadensis: 4(2):235
Antilles
Biomphalaria glabrata, Schistosoma,
S. mansoni: 4(1):120
Apache Lake, AZ
Corbicula fluminea, Ictiobus bubalus,
|. cyprinellus, |. niger: S2:7-39
Apalachee Bay, FL: 2:1-20
Apalachicola River, FL
Anodonta imbecilis: 4(2):231-232.
Corbicula fluminea: S2:7-39
Appomattox River, VA
Corbicula fluminea: S2:7-39
Arabian Gulf
Acanthopleura vaillantii, Chiton
(Acanthopleura) haddoni, C. lamyi, C.
peregrinus, C. (Rhyssoplax) affinis,
Notoplax (Notoplax) arabica:
6(1):115-130
Arabian Sea
Acanthopleura vaillantii, Callistochiton
adenensis, Chiton fosteri, C.
peregrinus, Ischnochiton yerbury,
Onithochiton erythraeus: 6(1):115-130.
Pupa affinis: 5(2):243-258. Tonicia
(Lucilina) sueziensis: 6(1):115-130
Arafura Sea
Thecacera pacifica: 5(2):243-258
Argentina
Corbicula fluminea: S2:1-5, 113-124.
C. leana: S2:113-124. Fissurellidea

megatrema, F. patagonica: 2:21-34.
Neocorbicula: S2:113-124. Trophon
geversianus: 3(1):11-26

Arizona (AZ)
Agua Fria River, Apache Lake, Col-
orado River: S2:7-39. Corbicula
fluminea: 4(1):81-88; S2:1-5, 7:39.
Gila River, Ictiobus bubalus, |.
cyprinellus, |. niger, Lake Martinez,
Salt River: S2:7-39. Sonorella:
4(1):113-114. Roosevelt Lake, Verde
River: S2:7-39

Arkansas (AR)
Arkansas River: 4(1):61-79, 115;
$2:1-5, 7-39, 193-201. Bayou Barthol-
omew, Black River, Bouef River:
$2:7-39. Buffalo National River: 1:97;
$2:193-201. Caddo River,
Chamagnoll Creek, Coon Bayou:
$2:7-39. Corbicula: S2:1-5, 59-61,
125-132. C. fluminea: 1:97; 4(1):61-79;
$2:7-39, 193-201. Dardanella Reser-
voir: S2:59-61. DeGray Lake:
$2:125-132. LaGrue Bayou,
LAnguille River, Little River,
Madison-Mariana Diversion Canal,
Manice Bayou, McKinney Bayou,
Ouachita River, Red River, Saline
River, St. Francis River, Spring
River, Strawberry River: S2:7-39.
Unionids, unspecified: 1:97. White
River: S2:7-39, 193-201

Arkansas River, AR, OK
Corbicula: S2:1-5. C. fluminea:
4(1):61-79; S2:7-39, 193-201.
Mollusca, unspecified, Paleontology:
4(1):115

Aruba
Acanthochitona andersoni, A. balesae,
A. rhodea, A. zebra, Arasji, Crypto-
conchus floridanus, Malmok, Rin-
con, Sero Colorado: 6(1):79-114

Atlantic Ocean
Onchidoris muricata, O. varians:
2:95. Opisthobranchia: 2:95-96.
Paziella: 3(1):11-26. Scaeurgus
unicirrhus: 6(2):207-211

Aucilla River, FL
Corbicula fluminea: S2:7-39

Australia
Amplirhagada: 1:98-99. Ascobulla
fischeri: 5(2):243-258. Avicennia:
4(1):112. Berthella pellucida:
5(2):197-214. Eucrassatella gibbosa:
2:83. Euselenops luniceps:
5(2):197-214. Kalinga ornata,
Kaloplocamus ramosus:
5(2):243-258. Littorina filosa, L.
scabra, Magnetic Island,
Metopograpsus: 4(1):112. Moreton
Bay: 5(2):197-214. Napier Range:
1:98-99. Nembrotha livingstonei:
5(2):243-258. New South Wales:
5(2):197-214; 6(1):115-130. Notobryon

wardi: 5(2):243-258.
Octopus tetricus: 6(1):45-48.
Onithochiton quercinus, O. rugulosus,
O. scholvieni: 6(1):115-130. Philinop-
sis cyanea, Placida dendritica:
5(2):243-258. Pleurobranchus peronii:
5(2):197-214. Polycera capensis, P
hedgpethi: 5(2):243-258.
Queensland: 2:57-61; 4(1):112;
5(2):197-214. Rhizophora: 4(1):112.
Roboastra gracilis: 5(2):243-258.
Thalamita crenata: 4(1):112. Thecacera
pennigera: 5(2):243-258. Trigonostoma
scalare: 2:57-61. Tylodina corticalis,
Umbraculum umbraculum:
5(2):197-214. Unionacea: 2:86-87.
Westaltrachia: 1:98-99.

Austria
Ancylus fluviatilis: 3(2):151-168

Avalon Bay, Trinidad
Acanthochiton balesae: 6(1):79-114

Bahamas
Abaco, Acanthochiles (Notoplax)
hemphilli, Acanthochitona andersoni,
A. balesae, A. lineata, A. pygmaea,
A. roseojugum, A. worsfoldi, A.
zebra, Acanthochitones spiculosus
astriger, Andros Island, Bahama
Beach Canal: 6(1):79-114. Boreo-
trophon aculeatus: 3(1):11-26.
Cancellaria reticulata: 4(1):113. Cat
Island, Choneplax lata, Chub Cay:
6(1):79-114. Crepidula navicula:
4(2):173-183. Cryptoconchus floridanus,
Dead Mans Reef, Elethura:
6(1):79-114. Fasciolaria tulipa:
4(1):113. Fernandez Bay, Fort Bay,
Gibson Cay, Great Exuma, Grand
Bahama, Green Turtle Cay, Harbour
Island, Isla Turramote, Long Island,
North Bimini, Salt Pond, Tamarind
Beach Reef: 6(1):79-114. Turbinella
angulata: 4(1):113. Utla Island:
6(1):79-114. Vexillum (Pusia) chick-
charneorum: 4(1):113. West Hawksbill
Creek: 6(1):79-114.

Bahama Beach Canal, Grand Bahama

Island

Acanthochiles (Notoplax) hemphilli,
Choneplax lata: 6(1):79-114

Bahrain
Acanthochitona woodwardi, Acantho-
pleura vaillantii, Ischnochiton yerbury,
Lepidozona luzonica, Notoplax
(Notoplax) arabica, Tonicia (Lucilina)
suezensis: 6(1):115-130

Baja California, Mexico
Ammonitellidae: 1:97. Biogeography:
1:97; 2:84-85. Bulimulidae: 1:97.
Cancellaria (Pyruclia) diadela, Cymia
chelonia: 2:84-85. Fissurellidea
bimaculata: 2:21-34. Haplotremati-
dae, Helminthoglyptidae: 1:97.
Holocene: 4(2):238-239. Melongena

256 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

melongena consors: 2:84-85.
Oreohelicidae: 1:97. Paleontology:
2:84-85; 4(2):238-239. Peninsula Ef-
fect: 1:97. Pliocene: 4(2):238-239.
Rapana bexoar vaquerosensis, R.
imperialis: 2:84-85. Rhabdotus,
Sonorella: 4(1):113-114. Speciation:
1:97. Solemya (Acharax) johnsoni:
$1:23-34. Spiracidae: 1:97. Tertiary,
Todos Santos, Turritella spp., T.
abrupta, Vasum pufferi: 2:84-85.

Baja California del Norte, Mexico
Arcticacea, Bernardina, B. margarita,
Bernardinidae, Cyamiacea, He/min-
thoglypta ayersiana, H. (Charodotes)
traskii: 3(1):103

Baja California Sur, Mexico
Amiantus sp., Anadara (Esmerarca)
sp., A. (Cunearca) nux: 4(1):12.
Arcticacea, Bernardina, B. bakeri,
Bernardinidae: 3(1):103. Calliostoma
hannibali, Calyptraea sp., Cardita
(Cardites) sp., Cerithium sp., Chione
(Chione) richthofeni, C. (Chionopsis)
sp., C. sp., Choromytilus pallio-
punctatus, Crassilabrum wittichi,
Crassispira starri, Crepidula sp.,
Crucibulum scutellatum: 4(1):1-12.
Cyamiacea: 3(1):103. Cyclinellas sp.,
Cymia heimi, Cypraea amandus,
Divalinga comis, Drillia (Clathrodrillia)
sp.: 4(1):1-12. Halodakra salmonea:
3(1):103. Hipponix pilosus, |sidro For-
mation, Knefastia sp., Lucina
(Lucinisca) sp., Macron hartmani,
Melongena melongena, M. melongena
consors, Mytilus canoasensis vidali,
Nassarius versicolor, Nerita
funiculata, Neverita (Glossaulax)
andersoni, Ostrea sp., Plicatula in-
ezana, Protothaca sp., Raeta sp.,
San Ignacio Formation,
Sanguinolaria toulai, Siphocypraea
henekeni, Siphonaria maura pica,
Solenosteira sp., Strombina sp.,
Tegula sp., Terebra burckhardati,
Theodoxus sp., Trachycardium sp.,
Trochita radians, T. spirata, T.
trochiformis, Turritella abrupta, T.
altilira, T. bifastigata, T. bosei, T.
costaricensis, T. crocus, T. inezana
bicarina, Vermetus contortus:
4(1):1:12

Barbados
Acanthochitona astriger, A. bonairen-
sis, A. rhodea, A. spiculosa, A.
worsfoldi, Acanthochitones
spiculosus astriger: 6(1):79-114.
Calliostoma apicinum, C. roseolum:
2:84

Barkley Lake, KY
Corbicula fluminea: S2:7-39

Barnegat Bay, NJ
Bankia gouldi: 4(1):89-99; S1:101-109.

Boveria teredinidi, B. zeukevitchi:
$1:101-109. Corbicula fluminea:
3(1):100-101. Crassostrea, Haplosporiad-
ium: $1:101-109. Mulinia lateralis:
4(1):39-42. Raritan River:
3(1):100-101. Teredo bartschi:
4(1):89-99; S1:101-109. T. furcifera:
$1:101-109. T navalis: 4(1):89-99;
$1:101-109

Barren Fork, Collins River, TN
Corbicula fluminea: S2:7-39. Lithasia
pinguis: 1:28

Barren River, KY
Pleurobema plenum: 1:28

Bay Champagne, LA
Thais haemastoma canaliculata:
6(2):189-197

Bay of Biscay
Hypselodoris cantabrica:
5(2):185-196. Tylodina perversa:
5(2):197-214

Bay of Fundy
Modiolus modiolus, Mulinia sp.,
Mytilus edulis, Placopecten
magellanicus: 4(1):104

Bayou Bartholomew, AR
Corbicula fluminea: S2:7-39

Bayou Cocodrie, LA
Corbicula fluminea: S2:7-39

Bayou Magasilla, LA
Corbicula fluminea: S2:7-39

Bayou Pierre, MS
Corbicula fluminea, Fusconaia flava,
Lampsilis ovata ventricosa, L. radiata
luteola, L. straminea claibornensis,
L. teres anodontoides, Leptodea
fragilis, Obovaria subrotunda,
Potamilus purpurata, Quadrula
pustulosa, Strophitus subvexus, Tox-
olasma texasensis, Tritogonia ver-
rucosa, Villosa lienosa: 4(1):21-23

Bayou Sorrel, LA
Corbicula fluminea: S2:7-39

Beach Fork Creek, WV
Corbicula fluminea: S2:7-39

Bear Creek, MS
Corbicula fluminea: S2:7-39

Bear Creek, TN
Corbicula fluminea: S2:7-39

Beaufort Inlet, NC
Chaetopleura apiculata, Diodora
cayenensis, Ischnochiton striolatus:
4(1):107-108

Belize
Acanthochiles (Notoplax) hemphilli,
Acanthochitona lineata, A. zebra:
6(1):79-114. Acochlidiacea: 2:95.
Ascobulla ulla, Berthellinia caribbea,
Bosellia mimetica: 5(2):259-280. Car-
rie Bow Cay, Choneplax lata:
6(1):79-114. Costasiella nonatoi, C.
ocellifera, Cyerce antillensis, Elysia
flava, E. papillosa, E. patina, E.
serca, E. sp., E. subornata, E. tuca,

Ercolania coerulea, E. funera,
Lobiger souverbiei, Oxynoe
antillarum, O. azuropunctata:
5(2):259-280. Pseudovermis: 2:95.
Tridachia crispata, Volvatella
bermudae: 5(2):259-280

Benbrook Lake, TX
Corbicula fluminea: S2:179-184

Bering Sea
Berryteuthis anonychus, B. magister,
Gonatus berryi, G. madokai, G. mia-
dendorffi, G. onyx, G. tinro: 2:89.
Nucella emarginata, Thais
emarginata: 1:105

Bermuda
Acanthochitona pygmaea: 6(1):79-114.
Ascobulla ulla: 5(2):259-280. Baileys
Bay: 6(1):79-114. Bosellia mimetica,
Costasiella nonatoi, C. occellifera,
Cyerce antillensis, C. crystallina,
Elysia flava, E. ornata, E. papillosa,
E. subornata, E. tuca, Lobiger
souverbiei, Oxynoe antillarum,
Placida sp., Volvatella bermudae:
5(2):259-280

Bethel Shoal, Key West, FL
Acanthochitona pygmaea: 6(1):79-114

Big Bigby Creek, TN
Corbicula fluminea: S2:7-39

Big Black Creek, MS
Corbicula fluminea: S2:7-39

Big Black River, MS
Corbicula fluminea: S2:7-39

Big Cedar Creek, AL
Corbicula fluminea: S2:7-39

Big Creek, MD
Corbicula fluminea: S2:7-39

Big Cypress National Preserve, FL
Liguus fasciatus aurantius, L.
fasciatus barbouri, L. fasciatus
castaneozonatus, L. fasciatus clenchi,
L. fasciatus elegans, L. fasciatus
floridanus, L. fasciatus livingstoni, L.
fasciatus lossomanicus, L. fasciatus
lucidovarius, L. fasciatus maiamien-
sis, L. fasciatus mosieri, L. fasciatus
ornatus, L. fasciatus roseatus, L.
fasciatus testudineus, L. fasciatus
walkeri: 5(2):153-157

Big Cypress River, TX
Corbicula fluminea: S2:7-39

Big Darby Creek, OH
Lasmigona costata: 2:82

Big Hickory Creek, TN
Corbicula fluminea: S2:7-39

Big Indian Creek, IN
Corbicula fluminea: S2:7-39

Big Moccasin Creek, VA
Alasmidonta viridis, Ambloplites
rupestris, Anadonta anatina,
Campostoma anomalum, Corbicula
fluminea, Cottus carolinae,
Etheostoma flabellare, E. rufilineaturm,
Fusconaia barnesiana, Lampsilis

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 257

fasciola: 5(1):1-7. Medionidus
conradicus: 5(1):1-7; 6(2):179-188.
Micropterus dolomieui, Nocomis
micropogon, Notropis coccogenis, N.
galacturus, Pisidium casertanum, P.
compressum: 5(1):1-7. Pleurobema
oviforme: 5(1):1-7; 6(2):179-188.
Sphaerium striatinum, Villosa
nebulosa: 5(1):1-7. V. vanuxemi:
5(1):1-7; 6(2):179-188

Big Nance Creek, AL
Corbicula fluminea: S2:7-39

Big River, MO
Corbicula fluminea: S2:7-39

Big Rock Creek, TN
Corbicula fluminea: S2:7-39

Big Seven Mile Creek, WV
Corbicula fluminea: S2:7-39

Big South Fork, Cumberland River, TN
Actinonaias pectorosa, Alasmidonta
atropurpurea, Elliptio crassidens, E.
dilatata, E. capsaeformis, Epioblasma
brevidens, Hemistena lata, Lampsilis
cardium, L. fasciola, L. ovata,
Lasmigona costata, Ligumia recta
latissima, Medionidus conradicus,
Pegias fabula, Pleurobema coccineum,
P. oviforme, Potamilus alatus, Ptycho-
branchus fasciolare, P subtentum,
Quadrula pustulosa, Strophitus un-
dulatus, Tritogonia verrucosa, Villosa
iris, V. taeniata, V. trabalis: 6(1):19-37

Big Swann Creek, TN
Corbicula fluminea: S2:7-39

Bimini
Acanthochitona pygmaea: 6(1):79-114

Bird Key Reef, FL
Acanthochitona roseojugum, Crypto-
conchus floridanus: 6(1):79-114

Biscayne Bay, FL
Codakia orbicularis: S2:23-34.
Granulina ovuliformis, Halodule
wrightii, Laurencia poitei: 4(2):185-199.
Lucina (Linga) pennsylvanica, Lucina
(Phacoides) pectinatus: $1:23-34.

Rissoina bryerea, South Biscayne Bay,

Thalassia testudinum, Tricolia

affinis affinis: 4(2):185-199
Biack River, AR

Corbicula fluminea: S2:7-39
Black River, MO

Corbicula fluminea: S2:7-39
Black Warrior River, AL

Corbicula fluminea: S2:7-39
Blanco River, TX

Corbicula fluminea: S2:7-39
Block Island, RI

Arctica islandica: S3:51-57
Blue River, TN

Corbicula fluminea: S2:7-39
Boeuf River, AR

Corbicula fluminea: S2:7-39
Bonaire

Acanthochitona andersoni, A. bon-

airensis, A. rhodea, Acanthochitones
spiculosus astriger, Choneplax lata,
Cryptoconchus floridanus: 6(1):79-114
Bonefish Key, FL
Acanthochitona balesae, A.
pygmaea, Cryptoconchus floridanus:
6(1):79-114
Boreal
Adamete viridula, Tritonium viridulum:
2:57-61
Borneo, Indonesia
Corbicula bitruncata, C. pullata:
$2:113-124
Bouge Phalia River, MS
Corbicula fluminea: S2:7-39
Bogue Sound, NC
Chaetopleura apiculata, Diodora
cayenensis: 4(1):107-108
Boreal
Adamete viridula, Tritonium viridulum:
2:57-61
Bourbeuse River, MO
Corbicula fluminea: S2:7-39
Bradley Creek, TX
Corbicula fluminea: S2:179-184
Bradley Reservoir, TX
Corbicula fluminea: S2:179-184
Brazil
Biomphalaria glabrata, B. straminea,
B. tenagophila: 1:67-70. Crepidula
protea: 1:110; 4(2):173-183. Croton
sp.-09: 1:67-70. Fissurellidea
megatrema: 2:21-34. Fusiturricula:
1:92. Littorina ziczac: 4(2):233.
Loligo sanpaulensis, Rio Grande do
Sol: 6(2):213-217. Siphonaria lessoni:
4(2):233. Turridae: 1:92
Brazos River, TX
Corbicula fluminea: S2:7-39, 179-164
British Columbia, Canada
Nucella emarginata: 1:105. Octopus
dofleini: 2:90. Thais emarginata:
1:105. Vancouver Island: 1:105; 2:90
Broad Creek, MD
Crassostrea virginica: S3:25-29
Brogley Rockshelter, WI
Actinonaias ligamentina carinaté,
Alasmidonta marginata, A. viridis
Amblema plicata, Anodonta grandis,
Anodontoides ferrussacianus, Arcidens
confragosus, Elliptio crassidens
crassidens, E. dilatata, E. dilatatus
delicatus, Fusconaia ebena, F. flava,
Lampsilis radiata luteola, L. teres
anodontoides, L. teres teres, L. ven-
tricosa, Lasmigona complanata, L.
compressa, L. costata, Ligumia recta,
Megalonaias nervosa, Potamilus
alatus, Quadrula pustulosa, Strophitus
undulatus undulatus, Venustaconcha
ellipsiformis ellipsiformis, Villosa iris
iris: 5(2):165-171
Brush Creek, OH
Corbicula fluminea: S2:7-39

Bryant Creek, MO
Corbicula fluminea: S2:7-39
Buck Creek, AL
Corbicula fluminea: S2:7-39
Buck Creek, KY
Corbicula fluminea: S2:7-39. Villosa
trabalis: 1:28
Buckatunna Creek, MS
Corbicula fluminea: S2:7-39
Buffalo National River, AR
Corbicula fluminea: 1:97; S2:7-39,
193-201. Unionids, unspecified: 1:97
Buffalo River, MS
Unionids: 4(1):21-23
Buffalo River, TN
Actinonaias ligamentina, A. pec-
torosa, Alasmidonta marginata, A.
viridus; 6(1):19-37. Corbicula
fluminea: S2:7-39. Cyclonaias tuber-
culata, Elliptio florentina walkeri,
Fusconaia barnesiana, F. barnesiana
bigbyensis, Hemistena lata, Lampsilis
cardium, L. fasciola, Lasmigona
complanata, L. costgata, Leptodea
fragilis, Lexingtonia dolabelloides
conradi, Obovaria subrotunda, O.
subrotunda lens, Pleurobema
oviforme, P. oviforme argenteum,
Potamilus ohioensis, Ptychobranchus
subtentum, Strophitus undulatus,
Toxolasma cylindrellus, Villosa iris, V.
taeniata, V. vanuxemensis: 6(1):19-37.
Burma
Ischnochiton (Ischnochiton) winck-
worthi: 6(1):115-130. Tricula spp.: 2:88
Burnt Corn Creek, AL
Corbicula fluminea: S2:7-39
Buttahatchie River, MS
Corbicula fluminea: S2:7-39
Buzzards Bay, MA
Chaetopleura apiculata: 6(1):69-78
Caballe Reservoir, NM
Corbicula fluminea: S2:7-39
Cabo Trough
Paleontology: 2:84-85
Caddo Creek, OK
Corbicula fluminea: S2:7-39
Caddo River, AR
Corbicula fluminea: S2:7-39
Cahaba River, AL
Corbicula fluminea: S2:7-39
Cahuma Lake, CA
Corbicula fluminea: S2:7-39
Calcasieu River, LA
Biogeography, Corbicula sp.: 2:86.
C. fluminea: S2:7-39. Sphaerium
spp., Unionids, unspecified: 2:86
California (CA)
Alamo Canal, All American Canal:
S2:7-39. Anaheim Bay, Los Angeles:
2:1-20; S2:7-39. Archidoris monterey-
ensis: 5(2):185-196. Argopecten
aequisulcatus: 4(2):241-242. Balanus
improvisus: $2:133-142. Berthellina

258 AMER. MALAC

engeli: 5(2):197-214. Boccardia
ligerica, Cahuma Lake: S2:7-39.
Cerithidea californica: 2:1-20.
Chaetogaster limnaei: S2:7-39. Chan-
nel Islands: 1:89; 2:83. Cimora
coneja: 5(2):287-292. Coachella
Water District, Colorado Aqueduct,
Colorado River, Columbia River:
S2:7-39. Cooper, James Graham:
1:89. Corbicula: S2:125-132. C.
fluminea: 4(1):81-88; S2:1-5, 7-39,
133-142. Corphium spinicoine, C.
stimpsoni: S2:7-39. Crepidula adunca:
3(1):33-40; 4(2):173-183. C. lingulata:
4(2):173-183. C. nummaria:
3(1):33-40. C. onyx: 1:110; 3(1):33-40;
4(2):173-183, 241-242. Crucibulum
spinosum: 4(2):241-242. Cryptom-
phalus (Helix) aspersa: 5(2):303-306.
Cuthona albocrusta: 5(2):287-292.
Delta-Mendota Canal: 4(1):81-88;
S2:7-39. Dyer Canal, El Capitan
Reservoir: S2:7-39. Eubranchus:
5(2):287-292. Eucrassinella fluctuata:
2:83. Evans Lake: S2:7-39. Fissurelli-
dea bimaculata: 2:21-34. Haliotis
cracherodii: 4(2):234-235. Halodakra,
H. (Halodakra) subtrigona, Helmintho-
glypta traskil: 3(1):103. Hermissenda
crassicornis: 4(2):205-216;
5(2):287-292. Ictalurus furcatus, Lake
Casitas, Lake Jennings, Lake Murray,
Lake Piru: S2:7-39. Lalia cockerelli:
5(2):287-292. Laevicardium
substriatum: 4(2):241-242. Livermore
Canal: S2:7-39. Loligo opalescens:
4(2):239. Macoma balthica,
Mayberry Cut, Merced River:
$2:7-39. Micrarionta opuntia: 3(1):98;
4(2):237. M. sodalis: 3(1):98;
4(2):237. Mitra idae: 1:91-92. Mohave
Desert: 1:89. Mokelumne Aqueduct,
Mokelumne River: S2:7-39. More-
teuthis pacifica. M. robusta: 4(2):241.
Mysella tumida: 4(2):234. Octopus:
4(2):234-235. O. bimaculatus: 2:90.
O. bimaculoides: 2:92-93;
4(2):241-242. Oligocene: 2:84-85.
Opuntia littoralis, Oreohelicidae:
2:98. Owens River: S2:7-39. Paleon-
tology: 3(1):98, 102-103. Phascolosoma
agassizii: 1:91-92. Physella virgata
virgata: 3(2):243-265. Placida den-
dritica: 5(2):243-258. Potatoee
Slough: S2:7-39. Radiocentrum
avalonense: 2:98. Rapana bexoar
vaquerosensis: 2:84-85. Russian
River: S2:7-39. Sacramento River:
4(1):81-88; S2:7-39, 125-132, 133-142.
Salinas River, Salton Sea: S2:7-39.
Salvia mellifera: 2:98. San Diego
City Water Works: S2:7-39. San
Francisco Bay: 2:1-20; S2:7-39. San

Jacinto Reservoir: S2:7:39. San
Joaquin River: 4(1):81-88; S2:7-39,
133-142. San Luis Reservoir:
$2:7-39. San Nicolas Island: 3(1):98;
4(2):237. Santa Barbara Channel:
5(2):287-292. Santa Catalina Island:
2:98. Saxidomus nutalli, Semele
decisa: 4(2):241-242. Shasta Lake:
S2:7-39. Sierra Nevada Mountains:
1:89. South Bay Aqueduct,
Stanislaus River, Stow Lake:
$2:7-39. Thunnus alaunga: 4(2):241.
Tolumne River: S2:7-39. Triopha
catalinae: 5(2):287-292. Xerarionta:
3(1):102-103

Caloosahatchee River, FL
Corbicula: S$2:125-132. C. fluminea:
$2:7-39

Caloosahatchian Province, FL
Paleontology: 2:79

Cambodia
Corbicula noetlingi, C. petiti:
$2:113-124

Campeche, Mexico
Acanthochitona pygmaea: 6(1):79-114

Canada
Alberta: 3(1):27-32. Amnicola limosa,
Anodonta grandis: 5(1):31-39. British
Columbia: 1:105; 2:90. Campeloma
decisum, Cincinnatia cincinnatiensis:
5(1):31-39. Cionella lubrica:
3(1):27-32. Crassostrea virginica:
$3:25-29. Elliptio complanata,
Gyraulus parvus, Helisoma anceps:
5(1):31-39. Illex illecebrosus: 2:51-56.
Lampsilis radiata: 5(1):31-39.
Macoma balthica, Manitoba: 1:90.
Melampus bidentatus: 4(1):121-122;
4(2):236. Musculium securis:
5(1):31-39. Neopanope sayi:
$3:59-70. New Brunswick:
4(1):121-122; 4(2):236; S3:59-70.
Newfoundland: 2:51-56. Nova Scotia:
$3:25-29. Nucella emarginata: 1:105.
Octopus dofleini: 2:90. Ontario,
Physella gyrina, Pisidium casertanum,
P compressum, P. ferrugineum, P
variable; 5(1):31-39. Prince Edward
Island: S3:25-29. Sphaerium rhom-
boideum, S. simile, S. striatinum:
5(1):31-39. Thais emarginata: 1:105.
Valvata tricarinata: 5(1):31-39. Van-
couver Island: 1:105; 2:90

Canary Islands
Discodoris fragilis: 5(2):243-258.
Hypselodoris webbi: 5(2):185-196.
Retusa truncata: 5(2):243-258

Cane Creek, MO
Corbicula fluminea: S2:7-39

Caney Fork River, TN
Actinonaias ligamentina gibba, A.
pectorosa, Alasmidonta autopur-
purea: 6(1):19-37. Amblema plicata:

. BULL. GEOGRAPHIC INDEX: 1983 - 1988

4(1):117. Cumberlandia monodonta:
6(1):19-37. Dromus dromas: 4(1):117.
Elliptio brevidens: 6(1):19-37. E. cras-
sidens, E. dilatata: 4(1):117. Epio-
blasma capsaeformis: 6(1):19-37. E.
florentina: 4(1):117. E. obliquata:
6(1):19-37. Fusconaias subrotunda,
Lampsilis ovata: 6(1):17-39. L. teres
teres: 4(1):117. Lasmigona complanata,
L. costata: 6(1):19-37. Ligumia recta,
Megalonaias nervosa: 4(1):117. Obli-
quaria reflexa, Pegias fabula, Pletho-
basus cicatricosus, Pleurobema gib-
berum: 6(1):19-37. Pleurobema plenum,
Pomtamilus alatus: 4(1):117.
4(1):117. Ptychobranchus subtentum,
Truncilla truncata, Villosa taeniata:
6(1):19-37

Cape Cod, MA
Panacca, P. arata, P fragilis:
3(1):103-104. Paleontology: 1:79

Cape Fear River, NC
Corbicula fluminea: S2:7-39. Elliptio
productus: 3(1):94

Cape Florida, FL
Cryptoconchus floridanus: 6(1):79-114

Cape Hatteras, NC
Paleontology: 2:79

Cape of Good Hope
Cancellaria lamellosa: 2:57-61.
Onithochiton wahlbergi: 6(1):115-130.
Opisthobranchia: 2:95-96

Caracas Baai, Curacao
Acanthochitona zebra: 6(1):79-114

Caribbean Sea
Aeolidiella alba, A. indica, Aplysia
dactylomela, A. juliana, Bertella
tupala: 5(2):243-258. Calliostoma
apicinum, C. pulchrum, C. roseolum:
2:84. Cancellaria reticulata: 2:57-61.
Chelidonura hirundinina:
5(2):243-258. Crassatella laevis: 2:83.
Cyphoma gibbosum: 2:84. Dolabrifera
dolabrifera, Lobiger souverbiei,
Micromelo undata: 5(2):243-258.
Mitridae: 3(1):97-98. Nudibranchia:
2:84. Octopus briareus, O. joubini:
6(1):45-48. Paziella: 3(1):11-26. Pleiop-
tygma, P. helenae: 3(1):97-98.
Polyplacophora: 1:91. Purpurella
patula: 4(1):110. Stylocheilus
longicauda, Umbraculum sinicum:
5(2):243-258. Voluta reticulata:
2:57-61. Volutidae: 3(1):97-98

Carrie Bow Cay, Belize
Acanthochiles (Notoplax) hemphilli,
Acanthochitona lineata, A. zebra,
Choneplax lata: 6(1):79-114

Cashie River, NC
Anodonta implicata, Lampsilis
ochracea, Ligumia nasuta:
3(1):104-105

Caspian Sea, USSR
Ancylus fluviatilis: 3(2):151-168

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 259

Cat Island, Bahamas
Acanthochiles (Notoplax) hemphilli,
Fernandez Bay: 6(1):79-114
Catawba River, NC
Corbicula: $2:125-132. C. fluminea:
$2:7-39
Cayman Islands
Acanthochiles (Notoplax) hemphilli,
Acanthochites spiculosus astriger,
Cryptoconchus floridanus, Grand
Cayman Island: 6(1):79-114.
Cayo Enrique, PR
Acanthochiles (Notoplax) hemphilli,
Acanthochitona lineata, A. pygmaea,
Cryptoconchus floridanus: 6(1):79-114
Cedar Creek, AL
Corbicula fluminea: S2:7-39
Cedar Creek Reservoir, TX
Corbicula fluminea: S2:179-184
Cedar Keys, FL
Acanthochitona pygmaea: 6(1):79-114
Cedar River, MI
Actinonaias ellipsiformis, Anodonta
grandis: 3(1):93. A. imbecilis: 3(1):93;
4(2):231-232. Anodontoides ferussac-
ianus, Fusconaia flava, Lampsilis
ovata, L. radiata, Lasmigona com-
pressa: 3(1):93
Celebes, Indonesia
Corbicula lindoensis, C. loehensis, C.
matanensis, C. planata: S2:113-124
Central America
Acanthochiles (Notoplax) hemphilli,
Acanthochiton balesae, Acantho-
chites rhodeus, Acanthochitona
andersoni, A. ferreirai: 6(1):79-114.
Acochlidiacea: 2:95. Aequipectin cir-
cularis: 4(1):119. Ascobulla ulla:
5(2):259-280. Atrina seminuda: 2:97.

Belize: 2:95; 5(2):259-280; 6(1):79-114.

Berthellinia caribbea, Bosellia
mimetica: 5(2):259-280. Costa Rica:
2:84; 3(1):98; 4(2):173-183.
Costasiella nonatoi, C. ocellifera:
5(2):259-280. Calyptraea conica, C.
mamillaris: 4(2):173-183. Cerithidea
montagnei, C. reevianum: 2:1-20.
Charonica tritonis: 2:84. Crepidula

cerithicola, C. convexa, C. dilatata, C.

echinus, C. fecunda, C. incurva, C.
lessoni, C. plana, C. striolata, Cruci-
bulum personatum, C. scutellatum,
C. spinosum, C. umbrella:
4(2):173-183. Cyerce antillensis:
5(2):259-280. Cypraea sp., C. talpa:
2:84. El Salvador: 2:97. Elysia flava,
E. papillosa, E. patina, E. serca, E.
sp., E. subornata, E. tuca, Ercolania
coerulea, E. funera: 5(2):259-280.
Favartia garretti: 2:84. Galeta Island:
6(1):79-114. Gatun Formation:

2:84-85; 4(1):1-12. Hipponix grayanus:

4(2):173-183. Honduras: 3(1):97-98;
6(1):79-114. Lobiger souverbiei:

5(2):259-280. Megapallifera: 4(2):238.
Mitridae, Nicaragua: 3(1):97-98.
Odostomia (Chrysallida): 4(1):122.
Ostrea irridescens: 4(1):119. Oxynoe
antillarum, O. azuropunctata:
5(2):259-280. Paleontology: 2:79,
84-85; 3(1):98. Pallifera: 4(2):238.
Panama: 2:1-20, 79, 84-85; 3(1):98;
4(1):1-12, 119, 122; 4(2):173-183;
6(1):79-114. Persicula pulchella: 2:84.
Philomycus: 4(2):238. Pinctada
mazatlanica: 4(1):119. Pinnidae: 2:97.
Pleioptygma, P. helenae: 3(1):97-98.
Protothaca asperimma: 4(1):119.
Pseudovermis: 2:95. Scalenostoma
subulata, Spondylus nicobarius:
2:84. Tridachia crispata:
5(2):259-280. Viriola abbotti: 2:84.
Volutidae: 3(1):97-98. Volvatella ber-
mudae: 5(2):259-280. Turridae:
3(1):98. Turritella abrupta: 4(1):1-12
Ceylon (Sri Lanka)
Cancellaria lamellosa, Trigonostoma
scalare: 2:57-61
Chaco River, Peru
Mollusca, unspecified: 3(1):96-97
Chain and Rocks Canal, IL
Corbicula fluminea: S2:7-39
Chamagnoll Creek, AR
Corbicula fluminea: S2:7-39
Channel Islands, CA
Cooper, James Graham: 1:89.
Eucrassinella fluctuata: 2:83
Charlotte Bay, FL
Acanthochitona pygmaea: 6(1):79-114
Chattahoochee River, GA, LA
Corbicula fluminea: S2:7-39
Chehalis River, WA
Corbicula fluminea: S2:7-39
Cherokee Starnes Site, TN
Archaeology, Actinonaias ligamentina,
Dromus dromus, Elliptio dilatata,
Epioblasma haysiana, Fusconaia
barnesiana, F. subrotunda, Lexingtonia
dolabelloides, Medionidus conraa-
icus, Pleurobema obliquum, Pleuro-
bema oviforme, Ptychobranchus
subtentum, Quadrula intermedia, A.
sparsa, Tellico River, Villosa iris:
3(1):41-44
Chesapeake Bay
Corbicula fluminea: S2:7-39.
Crassostrea virginica: 1:108; S3:5-10,
11-16, 17-23, 25-29. Haplosporidia
nelsoni: S3:5-10, 17-33. Paleontology:
2:79
Choptank River, MD
Crassostrea virginica: S3:25-29
Cibae Valley, Dominican Republic
Cercade Formation, Gurabo Forma-
tion, Mao Formation, Paleontology,
Turridae: 3(1):98
Chickahominy River, VA
Corbicula fluminea: S2:7-39

Chickamauga Creek, GA
Corbicula fluminea: S2:7-39
Chickasawhatchee River, GA
Corbicula fluminea: S2:7-39
Chickasawhay River, MS
Corbicula fluminea: S2:7-39
Chile
Buchanania onchidioides, Fissure-
lidae annulus, Fissurella patagonica,
Pupillaea annulus: 2:21-34. Trophon
geversianus: 3(1):11-26
China, Peoples Republic of (PRC)
Anodonta woodiana, Batissa
(Cyrenobatissa) subsulcata:
5(1):91-99. Biogeography: 2:88. Cor-
bicula aurea: S2:113-124. C.
fluminalis: 5(1):91-99; S2:113-124,
203-209. C. fluminea: 5(1):91-99;
$2:113-124, 203-209. C. /argillierti:
$2:113-124. C. manilensis: S2:1-5;
$2:113-124. C. nitens: S2:113-124.
Dali District, Dianchi Lake,
Jinghong, Kunming: 2:88. Lake
Hwama: S2:113-124. Lamprotula leai,
Limnoperna fortuei: 5(1):91-99.
Meghimatium: 4(2):238. Musculium
lacustre: 5(1):91-99. Parasitology:
2:88. Pearl River: S2:113-124,
203-209. Philomycidae: 4(2):238.
Polymesoda (Geloina) erosa:
5(1):91-99. Poyang Lake, San-Men-
Hsia Reservoir: S2:113-124. Tricula
spp.: 2:88. Triculinae: 3(1):96. Tung-
ting Lake: S2:113-124. Union
douglasiae: 5(1):91-99. Yangtze River,
Yellow River: S2:113-124
Chipola River, FL
Corbicula fluminea: S2:7-39
Choctawhatchee River, AL
Corbicula fluminea: S2:7-39
Choctawatchee River, FL
Corbicula fluminea: S2:7-39
Chowan River, NC
Corbicula fluminea: S2:219-222
Chub Cay, Bahama Islands
Acanthochites spiculosus astriger:
6(1):79-114
Chunky River, MS
Corbicula fluminea: S2:7-39
Cibao Valley, Dominican Republic
Cercado Formation, Gurabo Forma-
tion, Mao Formation, paleontology,
Turridae: 3(1):98
Clark Sound, SC
Mercenaria mercenaria: 4(2):149-155
Clear Fork, Trinity River, TX
Corbicula fluminea: S2:151-166
Clinch River, TN, VA
Actinonaias ligamentina: 4(1):25-37;
6(1):19-37. A. ligamentina gibba, A.
pectorosa, Alasmidonta marginata,
A. viridus: 6(1):19-37. Amblema
plicata: 4(1):25-37; 6(1):19-37.
Anodonta grandis grandis,

260 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

A. suborbiculata 6(1):19-37. Bivalvia,
unspecified: 4(2):231. Campeloma
sp., Conrdailla caelata: 4(1):25-37.
Corbicula fluminea: S2:7-39, 167-178,
Cumberlandia monodonta: 6(1):19-37.
Cyclonaias tuberculata: 4(1):25-37,
6(1):19-37. Cyprogenia irrorata:
4(1):25-37. C. stegaria: 4(1):25-37;
6(1):19-37. Dromus dromas:
4(1):25-37. D. dromas dromas, D.
dromas caperatus: 6(1):19-37. Elimia
sp.: 4(1):25-37. Ellipsaria lineolata:
6(1):19-37. Elliptio crassidens, E.
dilatata: 4(1):25-37; 6(1):19-37. E.
dilatata subgibbosus: 6(1):19-37.
Epioblasma arcaeformis: 4(1):25-37;
6(1):19-37. E. biemarginata: 6(1):19-37.
E. brevidens, E. capsaeformis:
4(1):25-37; 6(1):19-37. E. florentina:
6(1):19-37. E. haysiana: 4(1):25-37;
6(1):19-37. E. lenior, E. lewisi:
6(1):19-37. E. obliquata, E. propinqua,
E. stewartsoni: 4(1):25-37; 6(1):19-37.
E. torulosa: 4(1):25-37. E. torulosa
gubernaculum: 6(1):19-37. E. tri-
guetra: 4(1):25-37; 6(1):19-37. E.
turgidula: 6(1):19-37. Fusconaia
barnesiana: 4(1):25-37; 6(1):19-37. F
barnesiana bigbyensis, F. barnesiana
tumescens, F., F. cor analoga, F.
cuneolus appressa, F. cuneolus
cuneolus: 6(1):19-37. F. subrotunda:
4(1):25-37; 6(1):19-37. F. subrotunda
lesuerianus, F. subrotunda pilaris,
Hemistena lata: 6(1):19-37. lo fluvialis:
4(1):25-37. Lampsilis abrupta, L.
cardium: 6(1):19-37. L. fasciola:
4(1):25-37; 6(1):19-37. L. orbiculata:
4(1):25-38. L. ovata: 4(1):25-37;
6(1):19-37. L. virescens, Lasmigona
complanata, L. holstonia: 6(1):19-37.
Lemiox rimosa: 4(1):25-37; 6(1):19-37.
Leptodea fragilis, L. leptodon:
6(1):19-37. Leptoxis (Athearnia)
crassa, L. praerosa: 4(1):25-37. Lex-
ingtonia dolabelloides: 4(1):25-37;
6(1):19-37. Lexingtonia dolabelloides
conradi: 6(1):19-37. Ligumia recta:
4(1):25-37; 6(1):19-37. L. recta
latissima: 6(1):19-37. Lithasia ver-
rucosa: 4(1):25-37. Medionidus con-
radicus, Obliquaria reflexa, Obovaria
retusa: 6(1):19-37. O. subrotunda
lens: 4(1):25-37. O. subrotunda
subrotunda, O. subrotunda lavigata,
Plethobasus cicatricosus, P.
cooperianus, P cyphyus: 4(1):25-37;
6(1):19-37. Pleurobema catillus:
6(1):19-37. P. clava: 4(1):25-37;
6(1):19-37. P. coccineum: 6(1):19-37. P
cordatum: 4(1):25-37; 6(1):19-37. P
oviforme, P. oviforme argenteum, P.
oviforme holstonse: 6(1):19-37. P
plenum: 1:27-30; 6(1):19-37.

P. rubrum: 6(1):19-37. Pleurocera
canaliculatum, P canaliculatum un-
dulatum: 4(1):25-37. Potamilus alatus:
6(1):19-37. Ptychobranchus fasciolare,
P. subtentum: 4(1):25-37; 6(1):19-37.
Quadrula cylindrica: 4(1):25-27. Q.
cylindrica cylindrica, Q. cylindrica
Strigulata: 6(1):19-37. Q. intermedia,
Q. metanevra, Q. pustulosa: 4(1):25-37;
6(1):19-37. Q. sparsa, Strophitus un-
dilatus, Toxolasma lividus glans, T.
lividus lividus, T. parva, Truncilla trun-
cata: 6(1):19-37. Unionids, unspeci-
fied: 1:93-94. Villosa fabalis, V. iris,
V. perpurpurea: 6(1):19-37. V.
taeniata: 4(1):25-37. V. trabalis:
4(1):25-37; 6(1):29-37. V. vanuxemensis:
6(1):19-37. V. vanuxemi: 4(1):25-37

Coachella Valley Water District, CA
Corbicula fluminea: S2:7-39

Coahulla Creek, GA
Corbicula fluminea: S2:7-39

Coal River, KY
Corbicula fluminea: S2:7-39

Cocos Island, Costa Rica
Charonia tritonis, Cypraea sp., C.
talpa, Favartia garetti, Persicula
pulchella, Scalenostoma subulata,
Spondylus nicobaricus, Viriola
abbotti: 2:84

Coetivy Island
Tonicia (Lucilina) sueziensis:
6(1):115-130

Coldwater River, MS
Corbicula fluminea: S2:7-39

Coles Creek, MS
Toxolasma texasensis, Uniomerus
tetralasmus: 4(1):21-23

Collins River, TN
Corbicula fluminea: S2:7-39. Lithasia
pinguis: 1:27-30

Colombia
Acanthochiles (Notoplax) hemphilli,
Acanthochites rhodeus, Cabo la
Veda: 6(1):79-114. Crassostrea rhizo-
phorae: 1:35-42. Paleontology, Jur-
ritella abrupta: 4(1):1-12

Colorado (CO)
Pupilla blandi, P hebes, P.
muscorum, P. sonorana, P. sterkiana,
P syngenes: 1:99

Colorado Aqueduct, CA
Corbicula fluminea: S2:7-39

Colorado River, AZ, CA
Corbicula fluminea: S2:1-5, 7-39

Colorado River, TX
Corbicula: $2:125-132. C. fluminea:
$2:7-39

Columbia River, CA
Corbicula fluminea: S2:7-39

Columbia River, OR, WA
Corbicula fluminea: S2:7-39

Comoro Archipelago
Chiton (Chiton) fosteri: 6(1):115-130

Compano Bay, TX
Fossils, Molluscan Communities:
1:89

Conasauga River, GA, TN
Anodonta gradis corpulenta, A. im-
bellicus: 6(1):19-37. Corbicula
fluminea: S2:7-39. Elliptio arctata, E.
dilatata, Epioblasma metastriata,
Lampsilis altilis, L. clarkiana, L.
ornata, L. straminea claibornensis,
Lasmigona holstonia, Medionidus
acutissimus, M. conradicus,
Pleurobema aldrichianum, P. georg-
ianum, P. hanleyanum, P. johannis, P.
perovatum, P. rubellum, P.
troschelianum, Ptychobranchus
greeni, Strophitus connasaugaensis,
Toxolasma lividus glans, T. parva,
Villosa iris, V. lienosa, V. vanuxemen-
sis, V. vanuxemensis umbrans, V.
vibex: 6(1):19-37

Concho River, TX
Corbicula fluminea: S2:7-39

Conecuh River, AL
Corbicula fluminea, Graptemys
pulchra: S2:7-39

Congaree River, SC
Elliptio angustata: 1:95

Connecticut (CT)
Amnicola limosa, Campeloma
decisum, Cipangopaludina chinensis:
5(1):9-19. Crassostrea virginica:
$3:25-29. Crepidula convexa, C.
plana: 4(2):173-183. Ferrissia fragilis,
F. parallela, Gyraulus circumstriatus,
G. deflectus, G. parvus, Helisoma
anceps, H. campanulatum, H.
trivolvis, Laevapex fuscus: 5(1):9-19.
Long Island Sound: S3:25-29.
Lyrogyrus granum, L. pupoidea,
Micromentus dilatatus, Physa an-
cillaria, P heterostropha, Planorbula
armigera, Promenetus exacuous,
Pseudosuccinea columella,
Stagnicola elodes, Valvata tricarinata,
Viviparus georgianus: 5(1):9-19

Cook Islands
Mellanella sp., Rarotonga, Stichopus
chloronatus: 2:83

Coon Bayou, AR
Corbicula fluminea: S2:7-39

Cooper River, SC
Corbicula fluminea: S2:7-39

Coosa River, AL
Corbicula fluminea: S2:7-39

Corregidor, Philippines
Cancellaria lamellosa: 2:57-61

Costa Rica
Acanthochitona ferreirai, Acantho-
chitona rhodea: 6(1):79-114. Calyptraea
conica, C. mamillaris: 4(2):173-183.
Charonia tritonis: 2:84. Crucibulum
personatum, C. scutellatum, C.
spinosum, C. umbrella: 4(2):173-183.

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 261

Cypraea sp., C. talpa, Favartia gar-
retti: 2:84. Hipponix grayanus:
4(2):173-183. Paleontology: 3(1):98.
Persicula pulchella, Scalenostoma
subulata, Spondylus nicobaricus,
Viriola abbotti: 2:84. Turridae: 3(1):98

Coyner Springs, VA
Goniobasis proxima: 3(1):99-100

Crab Orchard Lake, IL
Corbicula fluminea: S2:7-39

Crawl Key, FL
Acanthochitona andersoni, A.
pygmaea, Cryptoconchus floridanus:
6(1):79-114

Cretaceous
Cerithiacea: 2:1-20

Croleavy Lough Outlet, Republic of

Ireland

Ancylus fluviatilis: 5(1):105-124

Cuba
Cerithidea scalariformis: 2:1-20.
Choneplax lata, Cryptoconchus
floridanus, Guantanimo Bay:
6(1):79-114. Oriente Province,
Polymita: 3(1):102-103

Cumberland River, KY, TN
Actinonaias ligamentina, A. ligamen-
tina gibba, A. pectorosa, Alasmidonta
autopurpurea, A. marginata, A.
viridus, Amblema plicata, A. plicata
perplicata, A. plicata plicata,
Anodonta grandis, A. imbecilis,
Anodontoides ferussacianus:
6(1):19-37. Corbicula fluminea:
4(1):81-88; S2:1-5, 7-39. Cumberlandia
monodonta, Cyclonaias tuberculata
tuberculata, C. tuberculata granifera,
Cyprogenia stegaria, Dromus dromas
dromas, Ellipsaria lineolata, Elliptio
crassidens, E. dilatata, Epioblasma
arcaeformis, E. brevidens, E. capsae-
formis, E. flexuosa, E. florentina, E.
florentina walkeri, E. haysiana, E.
lenior, E. obliquata, E. stewartsoni, E.
torulosa, E. torulosa torulosa, E. tri-
quetra, Fusconaia ebena, F. flava, F.
subrotunda, Hemistena lata, Lamp-
silis abrupta, L. cardium, L. fasciola,
L. ovata, L. teres anodontoides, L.
teres teres, Lasmigona complanata,
L. costata, Leptodea fragilis, Lexing-
tonia dolabelloides, Ligumia recta
latissima, Medionidus conradicus,
Megalonaias nervosa, Obliquaria
reflexa, Obovaria olivaria, O. retusa,
O. subrotunda, Pegias fabula, Pletho-
basus cicatricosus, P cooperianus,
P. cyphyus, P cyphyus compertus,
Pleurobema catillus, P clava, P coc-
cineum, P. cordatum, P gibberum, P
oviforme, P. plenum, P. rubrum,
Potamilus alatus, P ohioensis,
Ptychobranchus fasciolare, P subten-
tum, Quadrula cylindrica, Q. fragosa,

Q. metanevra, Q. pustulosa, Q.
quaarula, Simpsonaias ambigua,
Strophitus undulatus, Toxolasma
lividus glans, T. lividus lividus, T.
parva, Tritogonia verrucosa, Truncilla
donaciformis, T. truncata, Villosa iris,
V. lienosa, V. taeniata picta, V.
taeniata punctata, V. taeniata
taeniata: 6(1):19-37. V. trabalis:
1:27-30; 6(1):19-37. V. vanuxemensis:
6(1):19-37

Curagao
Acanthochites rhodeus, Acantho-
chitona andersoni, A. rhodea, A.
zebra, Acanthochitones spiculosus
astriger, Caracas Baai, Choneplax
lata, Piscadera Baai, Spaanse
Water: 6(1):79-114

Current River, MO
Cyclonaias tuberculata, Diversity,
Endangered Species, Fusconaia
ozarkensis, Lampsilis orbiculata, L.
reeviana, Pleurobema coccineum,
Ptychobranchus occidentalis, Villosa
iris iris: 2:85

Cushman Brook, MA
Margaritifera margaritifera: 4(1):13-19

Cuttyhunk Island, MA
Arctica islandica: S3:51-57

Cypress Creek, AL
Corbicula fluminea: S2:7-39

Cypress Creek Canal, FL
Corbicula fluminea: S2:7-39

Dallas Component, McMahan Site, TN
Actinonaias ligamentina, Alasmidonta
marginata, A. viridis, Amblema
plicata, Anodonta grandis,
Campeloma decisum, Cyclonaias
tuberculata, Cyprogenia stegaria,
Dromus dromas, Elliptio crassidens,
E. dilatata, Epioblasma arcaeformis,
E. brevidens, E. capsaeformis, E.
florentina, E. haysiana, E. steward-
soni, E. torulosa, Fusconaia
subrotunda, Hemistena lata, lo
fluvialis, Lampsilis fasciola, L. ovata,
Lasmigona costata, L. holstonia,
Lemiox rimosus, Leptoxis praerosa,
Lexingtonia dolabelloides, Ligumia
recta, Lithasia (Angitrema) verrucosa,
Medionidus conradicus, Obovaria
subrotunda, Plethobasus
cooperianus, P. cyphyus, Pleurobema
cordatum, P. oviforme, P plenum, P
rubrum, Pleurocera canaliculatum, P
parvum, Potamilus alatus, Ptycho-
branchus fasciolaris, P subtentum,
Quadrula cylindrica, Q. pustulosa, Q.
sparsa, Toxolasma lividus, Villosa
iris, V. trabalis: 6(2):165-178.

Damariscotta River, ME
Amnicola winkleyi, Cincinnatia wink-
leyi, Hydrobia truncata, Spurwinkia
Salsa: 4(1):101-102

Dardanelle Reservoir, AR
Corbicula: S2:59-61
Dauphin Island, AL
Corbicula fluminea: S2:7-39
Dead Mans Reef, Grand Bahama
Acanthochitona pygmaea: 6(1):79-114
DeGray Lake, AR
Corbicula: S2:125-132
Delaware (DE)
Corbicula fluminea: 4(1):81-88;
$2:7-39. Crassostrea virginica:
1:35-42. Nanticoke River: 4(1):81-88
Delaware River, NJ
Corbicula fluminea: S2:1-5, 7-39.
Elliptio productus: 3(1):94. Lim-
nodrilus spp., Peloscolex ferox, Pro-
cladius culiciformis, Sphaerium
transversum: S2:7-39
Delta-Mendota Canal, CA
Chaetogaster limnaei: S2:7-39. Cor-
bicula fluminea: S2:1-5, 7-39
Denmark
Lake Eorom, Pisidium subtruncatum:
5(1):41-48
Detroit River, MI
Actiononaias carinata, Alasmidonta
marginata, A. viridis, Amblema
plicata, Anodonta grandis grandis,
A. imbecilis, Anodontoides ferussaci-
anus, Caruncula parva, Cyclonaias
tuberculata, Dysnomia sulcata deli-
cata, Dysnomia torulosa rangiana,
Dysnomia triquetra, Elliptio dilatata,
Fusconaia flava, F. subrotunda,
Lampsilis fasciola, L. ovata, L. radi-
ata luteola, L. ventricosa, Lasmigona
complanata, L. compressa, L. costata,
Leptodea fragilis, L. leptodon,
Ligumia nasuta, L. recta, Obliquaria
reflexa, Obovaria olivaria, O.
subrotunda, Pleurobema coccineum,
Proptera alata, Ptychobranchus
fasciolare, Quadrula pustulosa, Q.
quaadrula, Simpsoniconcha ambigua,
Strophitus undulatus, Truncilla
donaciformis, T. truncata, Villosa
fabalis, V. iris: 3(1):105
Dianchi Lake, PRC
Tricula sp.: 2:88
Dix River, KY
Corbicula fluminea: S2:7-39
Dominican Republic
Acanthochitones spuculosus astriger:
6(1):79-114. Biomphalaria glabrata:
1:107. Cercado Formation, Cibao
Valley, Gurabo Formation, Mao For-
mation, Paleontology, Turridae:
3(1):98
Drivers Branch, AL
Corbicula fluminea: S2:7-39
Drunkeman’s Key, Jamaica
Acanthochiton balesae: 6(1):79-114
Dry Tortugas, FL
Acanthochiles (Notoplax) hemphillli,

262 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Acanthochitona andersoni, A. roseo-
jugum, A. zebra, Cryptoconchus
floridanus: 6(1):79-114

Duck Key, FL
Acanthochitona pygmaea: 6(1):79-114

Duck River, TN
Actinonaias ligamentina, A. pec-
torosa, Alasmidonta marginata, A.
viridus, Amblema plicata, Anodonta
grandis, A. imbecilis: 6(1):19-37. Cor-
bicula fluminea: S2:7-39. Cumberlan-
dia monodonta, Cyclonaias tuber-
culata, Cyprogenia stegaria, Ellipsaria
lineolata, Elliptio crassidens, E.
dilatata, Epioblasma brevidens, E.
capsaeformis, E. florentina, E. floren-
tina walkeri, E. lenior, E. lewisi, E.
torulosa, E. triquetra, E. turgida,
Fusconaia barnesiana, F. barnesiana
bigbyensis, Hemistena lata, Lampsilis
cardium, L. fasciola, L. ovata, L.
teres anodontoides, Lasmigona com-
planata, L. costata, L. holstonia,
Lemiox rimosus, Leptodea fragilis, L.
leptodon, Lexingtonia dolabelloides,
Lexingtonia dolabelloides conradi,
Ligumia recta latissima: 6(1):19-37.
Lithasia pinguis: 1:27-30. Medionidus
conradicus, Megalonaias nervosa,
Obliquaria reflexa, Obovaria retusa,
O. subrotunda, O. subrotunda lens,
Plethobasus cooperianus, P. catillus,
P cordatum, P. oviforme, P. oviforme
arghenteum, P. oviforme holstonse, P
rubrum, Potamilus alatus, P ohioen-
sis, Ptychobranchus fasciolare, P
subtentum, Quadrula cylindrica, Q.
fragosa, Q. intermedia, Q. pustulosa,
Q. quadrula, Strophitus undulatus,
Toxolasma cylindrellus, T. lividus
glans, Tritogonia verrucosa, Truncilla
donaciformis, T. truncata, Villosa
fabalis, V. iris, V. taeniata, V. vanux-
emensis: 6(1):19-37

Duplin Formation
Teinostoma nana: 4(1):39-42

Dutch Bay, Sri Lanka
Ischnochiton (lschnochiton) winck-
worthi: 6(1):115-130

Dyer Canal, CA
Corbicula fluminea: S2:7-39

Eagle Creek, KY
Corbicula fluminea: S2:7-39

Eagle Mountain Lake, TX
Aplocinotus grunniens, Corbicula
fluminea: S2:7-39

East Africa
Acteon fortis, Pleurobranchus brockii:
5(2):243-258

East Fork of Little Sandy River, KY
Lampsilis radiata luteola: 2:86

East Rock Creek, TN
Corbicula fluminea: S2:7-39

Easter Island
Julia zebra, Phanerophthalmus

smaragdinus, Smaragdinella
calyculata: 5(2):243-258

Ecuador
Angostura Formation: 4(1):1-12.
Crassatellinae: 2:83. Esmereldas
Formation: 2:84. Eucrassatella
digueti: 2:83. Mollusca, unspecified:
2:84. Paleontology: 3(1):98. Solemya
(Acharax) johnsoni: $1:23-34. Tur-
ridae: 3(1):98. Turritella abrupta, T. in-
ezana, T. ocoyana: 4(1):1-12

Eden River, NC
Corbicula fluminea: S2:7-39

Edisto River, SC
Corbicula fluminea: S2:7-39

El Capitan Reservoir, CA
Corbicula fluminea: S2:7-39

E! Salvador
Atrina seminuda, Pinnidae: 2:97

Elbow Reef, FL
Acanthochitona andersoni: 6(1):79-114

Elephant Butte Reservoir, NM
Corbicula fluminea: S2:7-39

Elethura, Bahamas
Acanthochiles (Notoplax) hemphilli:
6(1):79-114

Elizabeth River, VA
Crassostrea virginica: S3:31-36

Elk River, TN
Actinonaias carinata: 1:43-50. A.
ligamentina: 6(1):19-37. A. pectorosa:
1:43-50; 6(1):19-37. Alasmidonta
calceolus: 1:43-50. A. marginata:
1:43-50; 6(1):19-37. A. minor: 1:43-50.
A. viridus: 6(1):19-37. Amblema
costata: 1:43-50. A. plicata: 1:43-50;
6(1):19-37. Anculosa praerosa:
1:43-50. Anodonta grandis: 1:43-50;
6(1):19-37. Campeloma sp., Carun-
culina lividus, C. moesta, C. moesta
cylindrella, Conrdailla caelata:
1:43-50. Corbicula fluminea: S2:7-39.
C. manilensis: 1:43-50. Cyclonaias
tuberculata: 6(1):19-37. Dromus
dromas: 1:43-50; 6(1):19-37.
Dysnomia biemarginata, D. brevidens,
D. capsaeformis, D. florentina, D.
haysiana, D. torulosa, D. triquetra:
1:43-50. Ellipsaria lineolata:
6(1):19-37. Elliptio crassidens:
1:43-50; 6(1):19-73. E. dilatata:
6(1):19-37. E. dilatatus: 1:43-50.
Epioblasma biemarginata, E.
brevidens, E. capsaeformis, E. floren-
tina, E. haysiana, E. obliquata, E.
torulosa, E. triquetra, E. turgida:
6(1):19-37. Fusconaia barnesiana, F.
barnesiana bigbyensis: 1:43-50;
6(1):19-37. F. cor: 6(1):19-37. F.
cuneolus: 1:43-50; 6(1):19-37. F.
edgariana: 1:43-50. F subrotunda:
1:43-50; 6(1):19-37. Goniobasis la-
quetra: 1:43-50. Hemisetna lata:
6(1):19-37. lo verrucosa lima: 43-50.

Lampsilis anodontoides: 1:43-50. L.
cardium: 6(1):19-37. L. fasciola, L.
ovata: 1:43-50; 6(1):19-37. L. ovata
ventricosa: 1:43-50. L. teres teres:
6(1):19-37. Lasmigona complanata, L.
costata: 1:43-50; 6(1):19-37. Lastena
lata: 1:43-50. Lemiox rimosus:
6(1):19-37. Leptodea fragilis: 1:43-50;

6(1):19-37. Leptoxis praerosa: 1:43-50.

Lexingtonia dolabelloides: 1:43-50;
6(1):19-37. Lexingtonia dolabelloides
conradi, Lithasia verrucosa lima:
1:43-50. Medionidus conradicus:
1:43-50; 6(1):19-37. Megalonaias
gigantea: 1:43-50. M. nervosa:
6(1):19-37. Obliquaria reflexa,
Obovaria subrotunda, O. subrotunda
lens, Pegias fabula: 1:43-50;
6(1):19-37. Plagiola lineolata: 1:43-50.
Pleurobema cordatum, P. oviforme, P
oviforme argentium: 1:43-50;
6(1):19-37. Pleurocera canaliculatum:
1:43-50. Potamilus alatus: 6(1):19-37.
Proptera alata: 1:43-50. Ptycho-
branchus fasciolare: 6(1):19-37. P
fasciolaris: 1:43-50. P subtentum,
Quadrula cylindrica, Q. intermedia,
Q. metanevra, Q. pustulosa, Q.
quaoarula: 1:43-50; 6(1):19-37.
Strophitus rugosus: 1:43-50. S. un-
dulatus: 1:43-50; 6(1):19-37. Tox-
olasma cylindrellus, T. lividus glans:
6(1):19-37. Tritogonia verrucosa, Trun-
cilla donaciformis, T. truncata, Villosa
fabalis, V. iris: 1:43-50; 6(1):19-37. V.
nebulosa: 1:43-50. V. taeniata:
1:43-50; 6(1):19-37. V. vanuxemensis:
6(1):19-37. V. vanuxemi: 1:43-50.

Elk River, WV
Corbicula fluminea: S2:7-39

Elkhorn Creek, KY
Corbicula fluminea: S2:7-39

Elliott Key, FL
Acanthochitona andersoni, A. zebra:
6(1):79-114

Elm Fork, Trinity River, TX
Corbicula fluminea: S2:179-184

Emory River, TN
Amblema plicata: 6(1):19-37. Cor-
bicula fluminea: S2:7-39. Elliptio
crassidens, E. dilatata, E. turgidula,
Fusconaia barnesiana, F. cuneolus
cuneolus, Lampsilis cardium, L.
fasciola, L. virescens, Lasmigona
costata, Leptodea fragilis, Medioni-
dus conradicus, Pleurobema oviforme,
P holstonse, Potamilus alatus,
Ptychobranchus fasciolare, Quadrula
pustulosa, Toxolasma lividus glans,
T. lividus lividus, Villosa iris, V. per-
purpurea, V. vanuxemensis:
6(1):19-37

Enewetak, Marshall Islands
Akera soluta, Bornella anguilla,

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 263

Chromodoris geometrica, Elysia
livida, E. vatae, Flabellina, Halgerda
wasinensis, Marianina rosea,
Platydoris cruenta: 5(2):243-258.
Pleurehdera haraldi: 5(2):197-214

Escambia River, AL, FL

Corbicula fluminea: S2:7-39

Esmereldas Formation, Ecuador

Mollusca, unspecified: 2:84

Europe

Acanthochitona bonairensis, A. com-
munis: 5(1):79-114. Admetula evulsa:
2:57-61. Ancylus fluviatilis:

3(2):151-168. Archidoris pseudoargus:

5(2):185-196. Atagema gibba:
5(2):243-258. Bay of Biscay:
5(2):185-196, 197-214. Berthella
plumula: 5(2):243-258. Buccinum
evulsum: 2:57-61. Chaetogaster lim-
naei: 3(2):151-168. Chromodoris
krohni: 5(2):185-196. Doto coronata,
D. pinnatifida, Elysia viridis:
5(2):243-258. Eukiefferiella sp., Fer-
rissia wautieri, Glossiphonia com-
planata: 3(2):151-168. Goniodoris
castanea: 5(2):243-258. Gulf of
Genoa: 5(2):197-214. Halichondria
panicea, Hypselodoris bilineata. H.
cantabrica, H. gracilis, H. messinen-
sis, H. valenciennesi: 5(2):185-196.
Jorunna tormentosa: 5(2):185-196,
243-258. Limacia clavigera:
5(2):243-258. Mexichromis tricolor,
Microciona astrosanguinea:
5(2):185-196. Nitzschia
actinastroides: 3(2):151-168. Octopus
vulgaris: 2:92. Paleontology: 2:57-61.
Placida dendritica: 5(2):243-258.
Pleurobranchea meckelii:
5(2):197-214. Polycera faeroensis:
5(2):185-196. P. quadrilineata, Retusa
truncata: 5(2):243-258. Rostanga
rubra, Runcina coronata:
5(2):185-196. Tergipes tergipes,
Thecacera pennigera, Tritonia
nilsodhneri: 5(2):243-258. Tylodina
perversa: 5(2):197-214. Umbraculum
sinicum: 5(2):243-258. Unionacea:
2:86-87. Vorticella sp.: 3(2):151-168

Evans Lake, CA

Corbicula fluminea: S2:7-39

Falaika Island, Kuwait

Chiton (Chiton) peregrinus:
6(1):115-130

Falcon Reservoir, TX

Anodonta imbecilis henryiana, A.
grandis: 2:86. Corbicula fluminea:
2:86; S2:7-39. Cyrtonaias tampico-
ensis berlandieri, Disconaia
salinasensis, Lampsilis teres,
Megalonaias gigantea, Popenaias
popei, Quadrula apiculata, Tox-
olasma parvus, Unionmerus
tetralasmus manubius: 2:86

Fall Creek, TN

Corbicula fluminea: S2:7-39

Fanning Island

Atys cylindrica, Elysia marginata,
Pupa sulcata: 5(2):243-258

Farriers Pond, VA

Pisidium casertanum: 5(1):49-64

Fernandez Bay, Cat Island, Bahamas

Fiji

Acanthochiles (Notoplax) hemphilli:
6(1):79-114

Acochlidiacea: 2:95. Acteon flam-
meus, Chromodoris inopinata:
5(2):243-258. Gastrohedyle, Hedylop-
sis, Meiomenia, Meiopriapulus fijien-
sis, Nananu-i-ra Island, Paraganitus
ellynnae, Philinoglossa, Pseudover-
mis, Psuedunela, Viti Levu Island,
Yasawa Island: 5(2):281-286

Finiand

Anodonta piscinalis: 5(1):41-48. Lake

Varaslampi, Pisidium amnicum:
5(1):41-48. Pisidium casertanum, P
conventus: 5(1):21-30. Siilaisenpuro
River, Sphaerium corneum: 5(1):41-48

Flat Creek, TN

Corbicula fluminea: S2:7-39

Flint River, AL, GA

Corbicula fluminea, Lampsilis
anoaontoides floridensis, L. uniomi-
natus, Quincucina infucata: S2:7-39

Florida (FL)

Acanthochiles (Notoplax) hemphilli:
6(1):79-114. Acanthochitona andersoni,
A. astrigera, A. balesae, A. bonairen-
sis, A. communis, A. hemphilli, A. in-
terfissa: 1:91. A. pygmaea: 1:91;
6(1):79-114. A. rhodea: 1:91;
6(1):79-114. A. roseojugum:
6(1):79-114. A. spiculosa: 1:91. A.
zebra: 6(1):79-114. Acochlidiacea:
2:95. Alvania auberiana: 4(2):185-199.
Anclote Key: 6(1):79-114. Anodonta
imbecilis: 4(1):117; 4(2):231-232.
Anomia simplex: 2:41-50. Apalachee
Bay: 2:1-20. Apalachicola River:
4(2):231-232; S2:7-39. Aplacophora:
3(1):93-94; 4(1):107. Aplysiopsis zebra:
5(2):259-280. Argopecten gibbus:
2:41-50. Ascobulla ulla: 5(2):259-280.
Aucilla River: S2:7-39. Berthellinia
caribbea: 5(2):259-280. Bethel
Shoal: 6(1):79-114. Big Cypress Na-
tional Preserve: 5(2):153-157. Bird
Key: 6(1):79-114. Biscayne Bay:
$1:23-34. Bivalvia, unspecified:
3(1):93, 93-94. Bonefish Key:
6(1):79-114. Bosella marcusi, B.
mimetica: 5(2):259-280. Caecum
nitidum: 4(2):185-199. Caliphylla
mediterranea: 5(2):259-280.
Caloosahatchee River: S2:7-39,
125-132. Caloosahatchian Province:

2:79. Campeloma geniculum, C.
parthenum: 3(1):99. Cape Florida:
6(1):79-114. Caretta caretta: 3(1):93.
Caudofoveata: 4(1):107. Cedar Key:
6(1):79-114. Cerithidea costata, C.
scalariformis: 2:1-20. Charlotte Har-
bor: 6(1):79-114. Chione cancellata:
2:41-50. Chipola River: S2:7-39.
Codakia orbicularis: S1:23-34. Cor-
bicula: $2:125-132. C. fluminea:
$2:1-5, 7-39. Costasiella ocellifera:
5(2):259-280. Crassostrea virginica:
$3:25-29. Crawl Key: 6(1):79-115.
Crepidula aculeata: 4(2):173-183. C.
convexa: 1:110; 4(2):173-183. C. for-
nicata: 1:110. C. plana: 1:110;
4(2):173-183. Cryptoconchus
floridanus: 6(1):79-114. Cyerce an-
tillensis: 5(2):259-280. Cylichnella
canaliculata: 1:91. Cypress Creek:
S2:7-39. Duck Key, Dry Tortugas,
Elbow Reef, Elliot Key: 6(1):79-114.
Elliptio icterina: 1:95; 4(1):117. E. pro-
ductus: 3(1):94. Elysia, E. canguzua,
E. chlorotica, E. evelinae, E. ornata:
5(2):259-280. E. papillosa: 4(2):232.
E. serca: 5(2):259-280. E. subornata:
4(2):232; 5(2):259-280. E. tuca:
4(2):232; 5(2):259-280. Elysiidae:
4(2):232. Ercolania coerulea, E.
funera, E. fuscata, E. fuscovittata:
5(2):259-280. Escambia River, Ft.
Lauderdale Canal: S2:7-39. Ft.
Pierce: 5(2):259-280. Garden Key:
6(1):79-114. Gastropoda, unspecified:
3(1):93, 93-94. Geiger Key:
5(2):259-280. Geukensia demissa
demissa, G. demissa granosissima:
5(2):173-176. Granulina ovuliformis:
4(2):185-199. Graptacme calamus:
1:100. Grassy Key: 5(2):259-280;
6(1):79-114. Gulfport: 6(1):79-114.
Halodule wrightii: 4(2):185-199.
Hermanea cruciata: 5(2):259-280.
Holmes Creek: S2:7-39. Hutchinson
Island: 6(1):79-114. Ichetucknee
River, Indian Prairie Canal: S2:7-39.
Indian River: 2:1-20, 35-40. Indian
River Lagoon: 5(2):259-280. Key Bis-
cayne: 4(2):185-199. Key Largo:
4(2):185-199; 5(2):259-280. Key West:
6(1):79-114. Kissimmee River, Lake
Buena Vista, Lake Hippochee, Lake
Jackson, Lake Lucy, Lake
Okeechobee, Lake Oklawaha, Lake
Palatlakaha: S2:7-39. Lake Talquin:
1:95; 3(1):99; S2:7-39. Lake Tsala,
Lampsilis claibornensis: S2:7-39.
Laurencia obtusa, L. poitei:
4(2):185-199. Liguus fasciatus: 1:98;
3(1):1-10. L. fasciatus alternatus:
3(1):1-10. L. fasciatus aurantius:
3(1):1-10; 5(2):153-157. L. fasciatus
barbouri: 3(1):1-10; 5(2):153-157. L.

264 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

fasciatus beardi, L. fasciatus capen-
sis: 3(1):1-10. L. fasciatus castane-
Zonatus: 3(1):1-10; 5(2):153-157. L.
fasciatus castaneus, L. fasciatus
cingulatus; 3(1):1-10. L. fasciatus
clenchi: 3(1):1-10; 5(2):153-157. L.
fasciatus crassus, L. fasciatus
crenatus, L. fasciatus deckerti, L.
fasciatus delicatus, L. fasciatus
dohertyi, L. fasciatus dryas, L.
fasciatus eburneus: 3(1):1-10. L.
fasciatus elegans: 3(1):1-10;
5(2):153-157. L. fasciatus elliottensis,
L. fasciatus evergladenensis, L.
fasciatus farnumi; 3(1):1-10. L.
fasciatus floridanus: 3(1):1-10;
5(2):153-157. L. fasciatus framptoni,
L. fasciatus fuscoflamellus, L.
fasciatus gloriasylvaticus, L.
fasciatus graphicus, L. fasciatus
humesi, L. fasciatus innomillatus, L.
fasciatus kennethi, L. fasciatus
lignumvitae, L. fasciatus lineolatus:
3(1):1-10. L. fasciatus livingstoni:
3(1):1-10; 5(2):153-157. L. fasciatus
lossmanicus: 3(1):1-10; 5(2):153-157.
L. fasciatus lucidovarius, L. fasciatus
luteus, L. fasciatus margaretae, L.
fasciatus marmoratus, L. fasciatus
matecumbensis: 3(1):1-10. L.
fasciatus miamiensis: 3(1):1-10;
5(2):153-157. L. fasciatus mosieri:
3(1):1-10; 5(2):153-157. L. fasciatus
nebulosus: 3(1):1-10. L. fasciatus or-
natus: 3(1):1-10; 5(2):153-157. L.
fasciatus osmenti, L. fasciatus pic-
tus, L. fasciatus pseudopictus:
3(1):1-10. L. fasciatus roseatus:
3(1):1-10; 5(2):153-157. L. fasciatus

septentrionalis, L. fasciatus simpsoni,

L. fasciatus solida, L. fasciatus
solidulus, L. fasciatus solisocassus,
L. fasciatus splendidus, L. fasciatus
subcrenatus: 3(1):1-10. L. fasciatus

testudineus: 3(1):1-10; 5(2):153-157. L.

fasciatus vacaensis, L. fasciatus ver-
sicolor, L. fasciatus violafumosus, L.
fasciatus vonpaulseni: 3(1):1-10. L.
fasciatus walkeri: 3(1):1-10;
5(2):153-157. L. fasciatus wintei:
3(1):1-10. Lobiger souverbiei:
5(2):259-280. Long Key Reef:
6(1):79-114. Lower Matecumbe Key:
4(2):185-199; 6(1):79-114. Lucina
(Linga) pennsylvanica, L.
(Phacoides) pectinatus: $1:23-34.
Marginella aureocincta: 4(2):185-199.
Main Canal: S2:7-39. Mashta Island:
4(2):185-199. Mayakka River:
S2:7-39. Mercenaria mercenaria:
4(2):149-155. Middle River Canal:
$2:7-39. Missouri Key: 6(1):79-114.
Mosquitoe Creek: 4(2):231-232;
S2:7-39. Mourgona germaineae:

5(2):259-280. No Name Key:
6(1):79-114. North Mosquitoe Creek:
$2:7-39. Ochlocknee River: 3(1):99;
S2:7-39. Oklawaha River: S2:7-39.
Orthalicus floridensis, O. reses, O.
reses nesodryas: 2:98. Oxynoe an-
tillarum. O. azuropunctata:
5(2):259-280. Paleontology: 2:79;
4(1):107. Palm Beach Inlet:
6(1):79-114. Panacca, P arata, P
fragilis: 3(1):103-104. Paziella:
3(1):11-26. Peanut Island: 6(1):79-114.
Periploma margaritaceum: 2:35-40.
Placida, P. kingstoni: 5(2):259-280.
Pseudovermis: 2:95. Punta Rassa:
6(1):79-114. Rissoella caribaea, Ris-
soina bryerea: 4(2):185-199. Rocky
Creek, St. Johns River: S2:7-39. St.
Joseph Bay: 4(2):185-199; S2:7-39.
St. Lucie Inlet: 2:41-50. San Key:
6(1):79-114. Sanibel Island: 2:41-50;
6(1):79-114. St. Andrews Bay:
6(1):79-114. Santa Fe River: S2:7-39.
Sarasota Bay: 6(1):79-114.
Scaphopoda: 3(1):93-95. Sebastian
Inlet: 5(2):259-280. Sister Creek:
6(1):79-114. Sky Lake: S2:7-39.
Smaragdia viridis viridemaris:
4(2):185-199. Solenogastres: 4(1):107.
South Biscayne Bay: 4(2):185-199.
Spring Creek, Steinhatchee River:
S2:7-39. Strombus costatus, S.
(Tricornis) costatus, S. (Tricornis)
leidyi, S. (Tricornis) mayacensis:
4(1):108. Suwanee River: 3(1):99;
S2:7-39. Tampa Bay, Tennessee
Reef: 6(1):79-114. Thais haemastoma
canaliculata: 4(2):201-203. Thalassia
testudinum, Tricolia affinis affinis, T.
thalassicola: 4(2):185-199. Tridachia
crispata: 4(2):232; 5(2):259-280.
Vaca Key: 6(1):79-114. Villosa villosa:
1:95; 4(1):117. Virginia Key:
4(2):185-199. Waccassa River:
S2:7-39. Wekiva River: S2:1-5, 7-39.
Werstern Sambo Reef, West Sum-
merland Key: 6(1):79-114. Withla-
coochee River, Yellow River:
$2:7-39. Zebina browniana:
4(2):185-199

Florida Escarpment
Mytilids, Patellids, Trochids,
Vesicomyids: 3(1):95-96

Floyds Fork, KY
Corbicula fluminea: S2:7-39

Formosa
Chromodoris alderi: 5(2):243-258

Fort River, MA
Margaritifera margaritifera: 4(1):13-19

Fountain Creek, TN
Corbicula fluminea: S2:7-39

France
Acanthochitona bonairensis:
6(1):79-114. Corbicula fluminalis:

$2:113-124. Embletonia pulchra,
Heaylopsis spiculifera, Pontohedyle
milaschewitschii: 5(2):303-306.
Theba pisana: 1:104. Unela glana-
ulifera: 5(2):303-306
French Broad River, TN
Actinonaias ligamentina gibba,
Alasmidonta viridus, Amblema plicata,
Anodonta grandis corpulenta,
Cyclonaias tuberculata tuberculata,
Elliptio crassidens, E. dilatata, Epio-
blasma arcaeformis, E. capsaefor-
mis, E. florentina, E. turgidula,
Fusconaia barnesiana, F. barnesiana
bigbyensis, F. barnesiana tumescens,
F. subrotunda lesuerianus, F. sub-
rotunda pilaris, Lampsilis cardium, L.
fasciola, Lasmigona costata, L.
holstonia, Lexingtonia dolabelloides,
Ligumia recta, Pegias fabula,
Plethobasus cooperianus, P
cyphyus, P cyphyus compertus,
Pleurobema cordatum, P. oviforme, P
oviforme argenteum, P. oviforme hol-
stonse, P plenum, P. rubrum,
Potamilus alatus, Ptychobranchus
fasciolare, Quadrula pustulosa,
Strophitus undilatus, Toxolasma cylin-
drellus, T. lividus lividus, Villosa iris,
V. vanuxemensis: 6(1):19-37
French Polynesia
Moorea Island, Partula mooreana, P.
suturalis: 1:103-104. P taeniata: 1:104
Ft. Lauderdale Canal, FL
Corbicula fluminea: S2:7-39
Ft. Pierce, FL
Aplysiopsis zebra, Ascobulla ulla,
Bosellia mimetica, Caliphylla mediter-
ranea, Cyerce antillensis, Elysia
canguzua, E. ornata, E. sp., E.
subornata, E. tuca, Lobiger souver-
biei, Onynoe antillarum, Placida
kingstoni, P. sp.: 5(2):259-280
Galapagos Islands
Evolution: 2:85. Paziella: 3(1):11-26
Galapagos Rift
Aplacophora: S1:23-34. Calyptogena
magnifica, Mytilidae, Shell Microstruc-
ture, Shell Secretion: 1:101. Simrothiella:
$1:23-34. Vesicomyidae: 1:101
Galeta Island, Panama
Acanthochiton balesae: 6(1):79-114
Gannew Brook, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124
Gantt Lake, AL
Corbicula fluminea: S2:7-39
Garden Key, FL
Acanthochiles (Notoplax) hemphilli,
Acanthochitona andersoni, Crypto-
conchus floridanus: 6(1):79-114
Garrison River, TN
Corbicula fluminea: S2:7-39
Gasconade River, MO
Corbicula fluminea: S2:7-39

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 265

Gasper River, KY
Corbicula fluminea: S2:7-39

Gatun Formation, Panama
Paleontology: 4(1):1-12

Gatunian Province, Atlantic
Paleontology: 2:79

Gatunian Province, Pacific
Paleontology: 2:79, 84-85

Geiger Key, FL
Costasiella ocellifera, Cyerce antillen-
sis, Elysia papillosa, E. sp., E. subor-
nata, E. tuca, Ercolania funera,
Lobiger souverbiei, Mourgona ger-
maineae, Oxynoe azuropunctata,
Tridachia crispata: 5(2):259-280

Georgia (GA)
Altamaha River: 3(1):94, S2:1-5, 7-39.
Anodonta imbecilis: 4(2):231-232.
Cerithidea scalariformis: 2:1-20.
Chattahoochee River, Chickamauga
Creek, Chickasawhatchee River,
Coahulla Creek, Consauga River:
S2:7-39. Corbicula: S2:1-5. C.
fluminea: S2:1-5, 7-39. Crassostrea
virginica: $3:31-36. Elliptio shepar-
diana: 3(1):94. Flint River, Lake Alla-
toona, Lampsilis anodontoides
floridensis, L. uniominatus, Little Oc-
mulgee River: S2:7-39. Mercenaria
mercenaria: 4(2):149-155. Ocmulgee
River: 4(2):231-232; S2:7-39.
Ogeechee River, Ohoopee Creek,
Oostanula River, Potato Creek,
Pound Creek, Quincuncina infucata:
$2:7-39. Savannah River: S2:1-5,
7-39; S3:31-36. Towaliga River, With-
locoochee River: S2:7-39

Germany, Federal Republic of
Ancylus fluviatilis: 3(2):151-168

Ghana
Aplysia dactylomela, A. juliana,
Dolabrifera dolabrifera, Favorinus
ghanensis, Godiva quadricolor,
Hypselodoris tema, Pruttfolis
pselliotes, Thecacera pennigera:
5(2):243-258

Gibson Cay, Bahama Islands
Acanthochitona roseojugum:
6(1):79-114

Gila River, AZ
Corbicula fluminea: S2:7-39

Glen River, Republic of Ireland
Ancylus fluviatilus: 5(1):105-124

Glencullen River, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124

Glennaddragh River, Republic of Ireland
Ancylus fluviatilus: 5(1):105-124

Grand Bahama Island
Acanthochiles (Notoplax) hemphilli,
Acanthochitona andersoni, A.
balesae, A. worsfoldi, A. zebra,
Acanthochitones spiculosus astriger,
Choneplax lata, Cryptoconchus
floridanus, Long Island, Salt Pond,

Silver Cove Canal, Tamarind Beach
Reef: 6(1):79-114

Grand Cayman Island
Acanthochiles (Notoplax) hemphilli,
Acanthochitones spiculosus astriger:
6(1):79-114

Grant River, WI
Alasmidonta marginata, Anodonta
grandis corpulenta, Fusconaia flava,
Lampsilis radiata luteola, L. ven-
tricosa, Lasmigona complanata, L.
costata, Leptodea fragilis, Ligumia
recta, Potamilus alatus, Quadrula
quadrula, Q. verrucosa, Strophitus
undulatus undulatus, Venustaconcha
ellipsiformis ellipsiformis: 5(2):165-171

Grassy Creek, TN
Corbicula fluminea: S2:7-39

Grassy Key, FL
Acanthochitona pygmaea: 6(1):79-114.
Ascobulla ulla, Bosellia marcusi, B.
mimetica: 5(2):259-280. Cryptoconchus
floridanus: 6(1):79-114. Cyerce an-
tillensis, Elysia subornata, E. tuca,
Ercolania funera, Oxynoe antillarum,
Placida kingstoni, Tridachia crispata:
5(2):259-280

Grassy Lake, FL
Corbicula fluminea: S2:7-39

Great Exuma, Bahamas
Acanthochiles (Notoplax) hemphilli,
Cryptoconchus floridanus: 6(1):79-114

Great Miami River, OH
Corbicula fluminea: 3(1):94; S2:125-132

Great Wicomico River, VA
Crassostrea virginica, Haplosporidium
nelsoni: S3:17-23

Green Grotto Caves, Jamaica
Fossil Terrestrial Gastropoda: 1:99-100

Green River, KY
Actinonaias carinata, Alasmidonta
viridis, Amblema plicata, Anodonta
grandis: 1:29. Corbicula fluminea:
S2:7-39. Cyclonaias tuberculata,
Cyprogenia irrorata, Elliptio crassidens,
E. dilatata, Epioblasma triquetra,
Fusconaia flava, Lampsilis anodon-
toides, L. ovata, L. radiata siliquoidea,
Lasmigona costata, Leptodea fragilis,
Ligumia recta, Megalonaias
gigantea, Obliquaria reflexa,
Obovaria retusa, O. subrotunda,
Plagiola lineolata, Plethobasus
cyphyus, Pleurobema coccineum, P
cordatum, P. plenum, P
pyramidatum, Proptera alata,
Ptychobranchus fasciolare, Quadrula
metanevra, Q. nodulata, Q. pustulosa,
Q. quadrula, Tritogonia verrucosa,
Truncilla truncata, Villosa lienosa, V.
ortmanni: 1:29

Green Turtle Cay, Bahama Islands
Acanthochitona andersoni, Acantho-
chitona pygmaea: 6(1):79-114

Greenlick Creek, TN
Corbicula fluminea: S2:7-39
Guadeloupe
Boreotrophon lacunellus: 3(1):11-26
Guadelupe
Choneplax lata: 6(1):79-114
Guadelupe River, TX
Corbicula fluminea: S2:7-39, 179-184
Guantanimo Bay, Cuba
Choneplax lata: 6(1):79-114
Guayamas Basin
Aplacophora, Falcidens, Neomenia,
Thyasira: $1:23-34
Guinea
Voluta cancellata: 2:57-61
Gulf of Aden
Callistochiton adenensis: 6(1):115-130
Gulf of Alaska
Berryteuthis anonychus:
4(2):240-241. Gonatopsis borealis:
2:89-90. Gonatus middendorfi:
4(2):240-241. Ommastrephes bar-
trami: 2:89-90; 4(2):240-241.
Onychoteuthis borealijaponica: 2:89-90
Gulf of Aqaba
Ischnochiton (lschnochiton) yerburyi:
6(1):115-130
Gulf of California
Chromodoris annulata: 5(2):243-258.
Eucrassatella digueti, E. gibbosa:
2:83
Gulf of Geonoa, Italy
Pleurobranchaea meckelii:
5(2):197-214
Gulf of Maine, ME
Aeolidia papillosa, Catriona gymnota,
Coryphella gracilis, C. nobilis, C.
pellucida, C. salmonacea, C. verrilli,
C. verrucosa, Cuthona concinna,
Eubranchus tricolor, Facelina boston-
iensis, Metridium senile:
5(2):287-292. Placopecten
magellanicus: 6(1):1-8. Setoaeolis
pilata: 5(2):287-292
Gulf of Mexico
Octopus burryi: 2:92. O. joubini:
6(1):45-48
Gulf of Oman
Acanthopleura vaillantii, Chiton pere-
grinus, C. (Rhyssoplax) affinis,
Ischnochiton yerbury: 6(1):115-130
Gulf of St. Lawrence
Geukensia demissa demissa, G.
demissa granosissima: 5(2):173-176
Gulf of Suez
Chiton (Rhyssoplax) affinis, Onitho-
chiton erythraeus, Tonicia (Lucilina)
sueziensis: 6(1):115-130
Gulfport, FL
Acanthochitona pygmaea: 6(1):79-114
Guyandotte River, WV
Corbicula fluminea: S2:7-39
Halstead Bayou, MS
Polymesoda caroliniana: 6(2):199-206

266

Harbour Island, Elethura, Bahamas

Acanthochiles (Notoplax) hemphillli:
6(1):79-114

Harpeth River, TN

Actinonaias ligamentina, Alasmidonta
viridis: 6(1):19-37. Corbicula fluminea:
S2:7-39. Cyclonaias tuberculata,
Dromus dromas, Elliptio dilatata,
Epioblasma florentina, E. florentina
walkeri, E. obliquata, Fusconaia
flava, Lampsilis cardium, L. fasciola,
L. teres anodontoides, Lasmigona
complanata, L. costata, Ligumia rec-
ta latissima, Obovaria subrotunda,
Potamilus ohioensis, Ptychobranchus
subtentum, Quadrula fragosa, Q.
pustulosa, Strophitus undulatus, Tox-
olasma lividus lividus, Tritogenia ver-
rucosa, Truncilla donaciformis,
Villosa taeiniata picta, V. vanuxemen-
Sis: 6(1):19-37

Hartwell Reservoir, SC

Corbicula fluminea: S2:7-39

Hatchie River, TN

Amblema plicata, Anodonta grandis,
A. grandis corpulenta, A. imbecilis,
A. suborbiculata, Arcidens con-
fragosus: 6(1):19-37. Corbicula
fluminea: S2:7-39. Fusconaia ebena,
F. flava, Lampsilis cardium satura, L.
teres teres, L. teres anodontoides,
Lasmigona complanata, Leptodea
fragilis, Ligumia subrostrata, Mega-
lonaias nervosa, Obovaria jacksoniana,
Plectomaris dombeyanus, Pletho-
basus cyphus, Pleurobema cor-
datum, Potamilus ohiensis, P pur-
purata, Quadrula pustulosa, Q.
quaadrula, Strophitus undulatus, Toxo-
lasma parva, T. texasensis, Tritogonia
verrucosa, Truncilla truncata,
Uniomerus declivis, U. tetralasmus,
Villosa lienosa, V. vibex: 6(1):19-37.

Hawaii (HI)

Acanthochitona viridis: 6(1):79-114.
Achatina fulica: 2:98-99. Achatinelli-
dae: 4(1):112-113. Aplysia oculifera:
5(2):243-258. Aspidodiadema
hawaiiensis: 2:83. Barleeia:
4(2):232-233. Berthella tulapa,
Bertellina citrina, Berthellinia schlum-
bergeri, Bornella stellifer, Bullina
lineata: 5(2):243-258. Caecum sep-
timentum: 4(2):232-233. Caloria in-
dica, Ceratosoma cornigerum:
5(2):243-258. Cerithium placidum:
4(2):232-233. Chelidonura hirundinina:
5(2):243-258. Chondrocidaris
gigantea: 2:83. Chromodoris asper-
sa, C. marginata: 5(2):243-258. Cor-
bicula fluminea: S2:7-39. Crassostrea
virginica: S3:25-29. Dendrodoris
denisoni, D. nigra, Discodoris fragilis,
Doriopsis pecten, Elysia halimedae,

AMER. MALAC. BULL. GEOGRAPHIC INDEX

Embletonia gracilis: 5(2):243-258.
Euchelus gemmatus: 4(2):232-233.
Eugiandia rosea: 2:98-99. Eulimidae:
2:83. Euselenops luniceps, Favorinus
japonicus: 5(2):243-258. Gibbula
marmorea: 4(2):232-233. Gonaxis
kibweziensis, G. quadrilateralis:
2:98-99. Gymnodoris alba, G. bicolor,
G. okinawae, Hexabranchus
sanguineus, Hydatina albocincta,
Hypselodoris infucata, H. maridadi-
lus: 5(2):243-258. Joculator ridicula,
Julia exquisita, Kellia rosea, Kermia
aniani, Koloonella hawaiiensis, Lep-
tothyra rubricincta, Leptothyra
verruca, Lienardia balfreata,
Lophocochlias minutissimus, Maui:
4(2):232-233. Melibe pilosa,
Micromelo undata, Noumea
decussata, N. varians: 5(2):243-258.
Oahu: 2:83. Okadaia elegans:
5(2):243-258. Pelseneeria sp.: 2:83.
Phestilla melanobranchia, Phyllidia
varicosa, Phyllobranchillus orientalis,
Phyllodesmium serratum, Pleuro-
branchus peronii, Plocamopherus
maculatus: 5(2):243-258. Prionoci-
daris hawailiensis: 2:83. Pupa
tessellata: 5(2):243-258. Rissoina
ambigua: 4(2):232-233. Scaeurgus
patagiatus: 6(2):207-211. Schwartzi-
ella gracilis, Scissurella pseudo-
equatoria: 4(2):232-233. Tambja
morosa: 5(2):243-258. Tricolia
variabilis, Trivia exigua, Vanikoro
cancellata: 4(2):232-233. Vitreolina
sp.: 2:83

Hills Creek, TN

Lithasia pinguis: 1:28. Unionids,
Unspecified: 1:93-94

Hiwassee River, TN

Alasmidonta viridis, Elliptio
crassidens, Fusconaia barnesiana, F.
barnesiana bigbyensis, F. barnesiana
tumescens, Lasmigona holstonia,
Pleurobema oviforme, P. oviforme
holstonse, P. oviforme argenteum,
Tritogonia verrucosa, Villosa iris, V.
trabalis, V. vanuxemensis: 6(1):19-37

Hocking River, OH

Corbicula fluminea: S2:7-39

Holmes Creek, FL

Corbicula fluminea: S2:7-39

Holston River, TN

Actinonaias ligamentina, A. liga-
mentina gibba, A. pectorosa,
Alasmidonta ravenelina, A.
marginata, A. viridus, Amblema
plicata: 6(1):19-37. Corbicula fluminea:
$2:7-39. Cumberlandia monodonta,
Cyclonaias tuberculata tuberculata,
Cyprogenia stegaria, Dromus dromas
dromas, D. dromas caperatus, Ellip-
tio crassidens, E. dilatata, Epioblasma

: 1983 - 1988

arcaeformis, E. biemarginata, E.
brevidens, E. capsaeformis, E. floren-
tina walkeri, E. haysiana, E. lenior, E.
lewisi, E. obliquata, E. propinqua, E.
stewartsoni, E. torulosa gubernacu-
lum, E. triquetra, E. turgidula,
Fusconaia barnesiana, F. barnesiana
bigbyensis, F. barnesiana tumescens,
F. cor analoga, F. cuneolus ap-
pressa, F. cuneolus cuneolus, F.
subrotunda, F. subrotunda pilaris,
Hemistena lata, Lampsilis abrupta, L.
cardium, L. fasciola, L. ovata,
Lasmigona complanata, L. costata,
L. holstonia, Lemiox rimosa, Leptodea
fragilis, L. leptodon, Lexingtonia
dolabelloides conradi, Ligumia recta,
Medionidus conradicus, Obliquaria
reflexa, Obovaria retusa, O. subrotun-
da subrotunda, O. subrotunda
lavigata, Pegias fabula, Plethobasus
cicatricosus, P cooperianus, P
cyphyus, Pleurobema catillus, P. coc-
cineum, P. cordatum, P. oviforme,, P.
oviforme argenteum, P. oviforme
holstonse, P plenum, P. rubrum,
Potamilus alatus, Ptychobranchus
fasciolare, P subtentum, Quadrula
cylindrica cylindrica, Q. cylindrica
strigulata, Q. intermedia, Q.
metanevra, Q. pustulosa, Q. sparsa,
Strophitus undulatus, Toxolasma
lividus lividus, Truncilla truncata,
Villosa fabalis, V. iris, V. perpurpurea,
V. vanuxemensis: 6(1):19-37

Holston River, North Fork, VA

Actinonaias pectorosa, Alasmidonta
marginata, A. minor, Fusconaia
barnesiana, F. edgariana, Lampsilis
fasciola, L. ovata, Lasmigona
costata, Lexingtonia dolabelloides:
3(1):104. Medionidus conradicus,
Pleurobema oviforme: 3(1):104;
6(2):179-188. Ptychobranchus
fasciolaris, P subtentum, Toxolasma
lividus, Villosa nebulosa: 3(1):104, V.
vanuxemi: 3(1):104; 6(2):178-188

Homochitto River, MS

Anodonta imbecilis, Elliptio
crassidens, Fusconaia flava, Lamp-
silis claibornensis, L. radiata luteola,
Toxolasma texasensis, Uniomerus
declivus, Villosa lienosa: 4(1):21-23

Honduras

Acanthochiles (Notoplax) hemphilli,
Acanthochitona roseojugum,
Anthonys Key, Choneplax lata:
6(1):79-114. Mitrridae: 3(1):97-98. Oak
Ridge: 6(1):79-114. Pleioptygma, P.
helenae: 3(1):97-98. Roatan:
6(1):79-114. Volutidae: 3(1):97-98

Hong Kong

Anodonta woodiana: 5(1):91-99. Cor-
bicula fluminea: 5(1):91-99; S2:113-124.

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 267

Limnoperna fortuei, Musculium
lacustre: 5(1):91-99. Perna viridis:
4(2):233; 5(2):159-164. Pisidium an-
nandalei, P. clarkeanum, Polymesoda
(Geloina) erosa: 5(1):91-99
Hormuz Island, Iran
Acanthopleura vaillantii: 6(1):115-130
Horn Lake, TN
Strophitus undulatus: 6(1):19-37
Horse Creek, AL
Margaritifera margaritifera: 4(1):13-19
Hudson River Basin
Mollusca, unspecified: 4(1):119-120
Hughes River, WV
Corbicula fluminea: S2:7-39
Hutchinson Island, FL
Acanthochitona pygmaea: 6(1):79-114
ichetucknee River, FL
Corbicula fluminea: S2:7-39
Idaho (ID)
Corbicula fluminea, Snake River:
$2:7-39
Illinois (IL)
Chain and Rocks Canal: S2:7-39.
Corbicula fluminea: S2:7-39, 63-67.
Crab Orchard Lake: S2:7-39.
Cumberlandia monodonta: 4(1):13-19.
Fusconaia ebena: 5(2):177-179.
Hendersonia occulta: 1:99. Illinois
River, Kankakee River: S2:7-39.
Kaskasia River: S2:7-39, 63-67. Lake
Springfield: S2:7-39. Ohio River:
5(2):177-179; S2:7-39. Saline River,
Sangamon River: S2:7-39
Illinois River, IL
Corbicula fluminea: S2:7-39
India
Acanthochitona mahensis:
6(1):115-130. Cerithidea (Cerithdeop-
Silla): 2:1-20. Corbicula krishnaea, C.
regularis, C. striatella: S2:113-124.
Miocene: 2:1-20. Perna viridis:
5(2):159-164. Sclerodoris apiculata:
5(2):243-258. Tricula sp.: 2:88
Indian Creek
Corbicula fluminea: S2:7-39
Indian Ocean
Acanthochitona ashbyi, Acantho-
pleura vaillantii, Chiton salihafui:
6(1):115-130. Opisthobranchia:
2:95-96. Scaeurgus unicirrhus:
6(2):207-211
Indian Prairie Canal, FL
Corbicula fluminea: S2:7-39
Indian River, FL
Cerithidea scalariformis: 2:1-20
Indian River Inlet, FL
Periploma margaritaceum: 2:35-40
Indian River Lagoon, FL
Elysia canguzua, E. chlorotica, E.
evelinae, E. serca, Ercolania funera,
E. fuscata, E. fuscovittata, Hermaea
cruciata, Placida kingstoni:
5(2):259-280

Indiana (IN)
Big Indian Creek, Corbicula
fluminea: S2:7-39. Epioblasma samp-
soni: 1:28. Lymnaea elodes:
3(2):143-150; 6(1):9-17. Ohio River,
Salt Creek, Wabash River, White
River: S2:7-39

Indo-Pacific
Acteon fortis, Aeolidiella alba, A. in-
dica, Aplysia dactylomela, A. juliana,
Berthella tupala, Bethellina citrina:
5(2):243-258. Buccinum scalare:
2:57-61. Bulla ampulla: 5(2):243-258.
Cancellaria lamellosa, Delphinula
trigonostoma: 2:57-61. Discodoris
fragilis, Dolabrifera dolabrifera,
Doriopsis pecten, Euselenops
luniceps, Halgerda formosa, H.
punctata, H. wasinensis, Lobiger
souverbiei, Oxynoe viridis, Pleuro-
branchella nicobarica, Pleurobranchus
brockii, P inhacae, P xhosa, Pupa
affinis, P. solidula: 5(2):243-258.
Scalptia nassa: 2:57-61. Stylocheilus
longicauda: 5(2):243-258. Trigona
pellucida: 2:57-61. Umbraculum um-
braculum: 5(2):243-258. Voluta
nassa: 2:57-61

Indonesia
Anodonta woodiana: 5(1):91-99.
Borneo, Celebes: S2:113-124.
Cerithidea, C. (Cerithdeopsilla), C.
rhizophorarum: 2:1-20. Corbicula
australis, C. bitruncata, C. gusta-
viana, C. javanica, C. lindoensis, C.
loehensis, C. matanensis, C.
moltkiana, C. planata, C. pulchella,
C. pullata, C. rivalis, C. sumatrana,
C. tobae, C. tumida: S2:113-124.
Java: 2:1-20; S2:113-124. Pliocene:
2:1-20. Sumatra: 2:1-20; S2:113-124.
Timor: S2:113-124

Inhaca Island
Onithochiton litteratus: 6(1):115-130

Intercoastal Waterway, SC
Corbicula fluminea: S2:7-39

lowa (IA)
Anodonta grandis grandis: 1:71-74.
Corbicula fluminea: S2:7-39. Cumber-
landia mondonta: 4(1):13-19. Hender-
sonia occulta: 1:99. Leptodea fragilis:
1:71-74. Mississippi River: S2:7-39

Iran
Acanthopleura vaillantii, Hormuz
Island: 6(1):115-130

Isidro Formation, Baja California Sur,

Mexico
Anadara (Cunearca) nux, Calyptraea
sp., Cerithium sp., Chione sp., Hip-
ponix pilosus, Melongena melongena,
Ostrea sp., Plicatula inezana, Pro-
tothaca sp., Siphocypraea henekeni,
Siphonaria maura pica, Tegula sp.,
Theodoxus sp., Trochita radians,

T. spirata, T. trochiformis, Turritella
altilira, T. crocus, Vermetus contor-
tus: 4(1):1-12

Isla Margarita, Venezuela
Acanthochitona venezuelana:
6(1):79-114

Isla Mujeres, Mexico
Acanthochitona pygmaea: 6(1):79-114

Isla Turramote, PR
Acanthochitona pygmaea, A. zebra:
6(1):79-114

Israel
Chiton huluensis: 6(1):115-130. Cor-
bicula fluminalis, Sea of Galilee:
$2:113-124. Ischnochiton (Ischno-
chiton) yerburyi: 6(1):115-130

Italy
Cepaea nemoralis, C. nemoralis
nemoralis, C. vindobonensis:
1:107-108. Gulf of Geonoa:
5(2):197-214. Littorina saxatilis:
1:92-93. Pleurobranchaea meckelii:
5(2):197-214

Ireland, Northern, UK
Embletonia pulchra: 5(2):303-306

Ireland, Republic of
Aille River: 5(1):105-124. Ancylus
fluviatilis: 3(2):151-168; 5(1):105-124.
Croleavy Lough Outlet, Gannew
Brook, Glen River, Glencullen River,
Glennaddragh River, Little Brosna
River, Lough Inch, Nore River, Owen
Doher River, Owenwee River, River
Liffey, Woodford River: 5(1):105-124

Jacks Ford River, MO
Diversity, Fusconaia ozarkensis,
Lampsilis reeviana, Ptychobranchus
occidentalis, Villosa iris iris: 2:85

Jalisco, Mexico
Bernardina margarita: 3(1):103

Jamaica
Acanthochiles (Notoplax) hemphili,
Acanthochitona balesae: 6(1):79-114.
Camaenidae: 3(1):102-103. Cryp-
toconchus floridanus, Drunkeman’s
Key: 6(1):79-114. Fossil Gastropoda,
Terrestrial, Green Grotto Caves:
1:99-100. Orthalicus undatus
jamaicensis: 2:98. Paleontology:
1:99-100; 3(1):98, 102-103.
Pleurodonte: 3(1):102-103. Turridae:
3(1):98

James River, VA
Corbicula fluminea: S2:7-39.
Crassostrea virginica: S3:17-23, 31-36.
Elliptio fisherianus, E. lanceolata, E.
productus: 3(1):94. Haplosporidium
nelsoni: S3:17-23

Japan
Cerithidea (Cerithdeopsilla): 2:1-20.
Chelidoneura fulvipunctata:
5(2):243-258. Corbicula felnouilliana,
C. fluminea, C. fluviatilis: S2:113-124.
C. japonica: $2:1-5, 113-124.

268

Java

Java

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

C. leana: $2:113-124, 203-209. C.
sandai: S2:1-5, 113-124. Cuthona an-
nulata, C. ornata, Goniodoris
castanea, Gymnodoris inornata,
Hydatina zonata: 5(2):243-258.
Meghimatium, Philomycidae:
4(2):238. Marioniopsis cyanobranch-
iata: 5(2):243-258. Miocene: 2:1-20.
Nembrotha lineolata, Noumea pur-
purea: 5(2):243-258. Perna viridis:
5(2):159-164. Placida dendritica:
5(2):243-258. Pliocene: 2:1-20.
Roboastra luteolata, Stiliger ornatus,
Thecacera pennigera: 5(2):243-258.
Unionacea: 2:86-87

, Indonesia

Cerithidea, C. rhizophorarum: 2:1-20.
Corbicula javanica, C. pulchella, C.
rivalis: S2:113-124. Pliocene: 2:1-20
Sea

Lepidozona (Lepidozona) luzonicus:
6(1):115-130

Jeffrey’s Basin, ME

Placopecten magellanicus: 6(1):1-8

Jericho Bay, ME

Placopecten magellanicus: 6(1):1-8

John Day River, OR

Corbicula fluminea: S2:7-39

Johnson Creek, TX

Corbicula fluminea: S2:7-39

Juan de Fuca Vent

Aplacophora, Neomenia, Simrothiella:

$1:23-34

Kanawha River, WV

Actinonaias lineola carinata,
Amblema plicata plicata, Anodonta
grandis grandis, A. imbecilis, Cor-
bicula fluminea, Cyclonaias tuber-
culata, Cyprogrenia stegaria, Ellip-
saria lineolata, Elliptio crassidens
crassidens, E. dilatata, Endangered
Species, Fusconaia maculata
maculata, Lampsilis fasciola, L.
obriculata, L. ovata, L. radiata
luteola, L. ventricosa, Lasmigona
costata, L. subviridis, Leptodea
fragilis, Ligumia recta, Megalonaias
nervosa, Obliquaria reflexa, Obovaria
subrotunda, Plethobasus cyphus,
Pleurobema cordatumx, P rubrum, P
sintoxia, Potamilus alatus,
Ptychobranchus fasciolaris, Quadrula
pustulosa pustulosa, Simpsonaias
ambigua, Strophitus undulatus un-
dulatus, Tritogonia verrucosa, Trun-
cilla truncata, Villosa iris iris:
2:85-86

Kankakee River, IL

Corbicula fluminea: S2:7-39

Kansas (KA)

Allogona profunda, Anguispira alter-
nata, A. kochi, Cepaea hortensis, C.
nemoralis, Helix aspersa, H.
pomacea, Mesodon clausus,

M. elevatus, M. thyroidus, Succinea
ovalis, Triodopsis albolabris alleni, T.
multilineata: 1:97-98

Kaskasia River, IL

Corbicula fluminea: S2:7-39; 63-67

Kentucky (KY)

Actinonaias carinata: 1:29. A.
ligamentina carinata: 1:31-34.
Alasmidonta viridis: 1:29. Amblema
plicata: 1:29, 31-34; 3(1):47-53. A.
plicata plicata: 3(1):47-53. Anodonta
grandis: 1:29. A. grandis grandis:
3(1):47-53. Archaeology: 1:31-34.
Buck Creek, Coal River: S2:7-39.
Corbicula: S2:125-132. C. fluminea:
3(1):47-53; S2:7-39. Cumberland
River: S2:7-39. Cyclonaias tuber-
culata, Cyprogenia irrorata: 1:29. C.
Stegaria: 1:31-34. Dix River, Eagle
Creek: S2:7-39. East Fork of Little
Sandy River: 2:86. Elimina sp.:
1:31-34. Elkhorn Creek: S2:7-39.
Elliptio crassidens: 1:29. E.
crassidens crassidens: 3(1):47-53. E.
Cilatata: 1:29, 1:31-34; 3(1):47-53.

Epioblasma sampsoni: 1:31-34. E. tri-

quetra: 1:29; 3(1):47-53. Floyds Fork:
$2:7-39. Fort Ancient People:
1:31-34. Fusconaia flava: 1:29,
1:31-34; 3(1):47-53. F maculata
maculata: 1:31-34. Gasper River:
S2:7-39. Goniobasis sp.: 1:31-34.
Green River, Kentucky Reservoir:
S2:7-39. Kentucky River, KY: 1:29,
31-34; S2:7-39. Kinniconick Creek:
3(1):47-53. Lampsilis anodontoides:
1:29. L. fasciola: 3(1):47-53. L. ovata:
1:29. L. radiata luteola: 2:86;
3(1):47-53. L. radiata siliquoidea:
1:29. L. ventricosa: 1:31-34;
3(1):47-53. Lasmigona costata: 1:29;
3(1):47-53. Leptodea fragilis: 1:29;
3(1):47-53. Licking River: S2:7-39.

Ligumia recta: 1:29. Lithasia obovata:

1:31-34. Little River: S2:7-39,
125-132. Magnonaias nervosa:
1:31-34. Megalonaias gigantea: 1:29.
Mississippi River, Mud River, Nolin
River: S2:7-39. Obliquaria reflexa:

1:29. Obovaria retusa, O. subrotunda:

1:31-34. Ohio River: S2:7-39. Pauzar
Rockshelter, Physa sp.: 1:31-34.
Plagiola lineolata: 1:29. Pleurobema
Clava: 1:31-34. P cordatum: 1:29,
1:31-34. P plenum: 1:29, 1:31-34. P
rubrum, P. sintoxia, Pleurocera
canaliculatum: 1:31-34. Potamilus
alatus: 3(1):47-53. Proptera alata,
Ptychobranchus fasciolare: 1:29. P
fasciolaris: 1:31-34; 3(1):47-53.
Quaadrula nodulata: 1:29. Q.
pustulosa: 1:29, 1:31-34. Q.
pustulosa pustulosa: 3(1):47-53. Q.
quadrula: 1:29, 1:31-34. Red River,

Rockcastle River, Salt River, Silver
Creek: S2:7-39. Simpsonaias ambi-
gua: 3(1):47-53. Slate Creek: S2:7-39.
Strophitus undulatus undulatus:
3(1):47-53. Tennessee River,
Tradewater River: S2:7-39. Tritogonia
verrucosa: 1:29; 3(1):47-53. Truncilla
truncata: 1:29. Tygarts Creek:
S2:7-39. Villosa iris iris, V. lienosa:
3(1):47-53

Kentucky Reservoir, KY, TN

Corbicula fluminea: S2:7-39

Kentucky River, KY

Actiononaias ligamentina carinata:
1:31-34. Amblema plicata: 1:29,
1:31-34. Archaeology: 1:31-34. Cor-
bicula flumina: S2:7-39. Cyprogenia
stegaria, Elimina sp., Elliptio dilatata,
Epioblasma sampsoni, Fort Ancient
People, Fusconaia flava, F. maculata
maculata, Goniobasis sp., Kentucky,
Lampsilis ventricosa, Lithasia
obovata, Magnonaias nervosa,
Obovaria retusa, O. subrotunda,
Pauzar Rockshelter, Physa sp.,
Pleurobema clava, P cordatum, P
plenum, P. rubrum, P. sintoxia,
Pleurocera canaliculatum, Ptycho-
branchus fasciolaris, Quadrula
pustulosa, Q. quadrula: 1:31-34

Kenya

Chiton (Chiton) fosteri: 6(1):115-130

Key Biscayne, FL

Alvania auberiana, Caecum nitidum,
Granulina ovuliformis, Halodule
wrighti, Laurencia poitei, Marginella
aureocincta, Rissoella caribaea, Ris-
soina bryerea, Smaragdia viridis
viridemaris, Thalassia testudinum,
Tricolia affinis affinis, T. thalassicola,
Zebina browniana: 4(2):185-199

Key Largo, FL

Alvania auberiana: 4(2):185-199.
Ascobulla ulla, Berthellinia caribbea,
Bosellia mimetica, Costasiella
ocellifera: 5(2):259-280. Crypto-
conchus floridanus: 6(1):79-114.
Cyerce antillensis, Elysia, E.
papillosa, E. patina, E. serca, E.
subornata, E. tuca, Ercolania
coerulea, E. funera, E. fuscata:
5(2):259-280. Halodule wrightii:
4(2):185-199. Hermaea cruciata:
5(2):259-280. Laurencia poitei:
4(2):185-199. Lobiger souverbiei,
Oxynoe antillarum, O. azuropunc-
tatum: 5(2):259-280. Thalassia
testudinum, Tricolia affinis affinis:
4(2):185-199. Tridachis crispata:
5(2):259-280

Key West, FL

Acanthochiles (Notoplax) hemphillli,
Acanthochitona andersoni, A.
pygmaea, Cryptoconchus floridanus:

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 269

6(1):79-114

Kinniconick Creek, KY
Amblema plicata plicata, Anondonta
grandis grandis, Corbicula fluminea,
Elliptio dilatata, E. crassideus
crassideus, Epioblasma triquetra,
Fusconaia flava, Lampsilis fasciola,
L. radiata luteola, L. ventricosa,
Lasmigona costata, Leptodea fragilis,
Potamilus alatus, Ptychobranchus
fasciolaris, Quadrula pustulosa
pustulosa, Simpsonaias ambigua,
Strophitus undulatus undulatus,
Tritogonia verrucosa, Villosa iris iris,
V. lienosa: 3(1):47-53

Kissimmee River, FL
Corbicula fluminea: S2:7-39

Korea
Corbicula colorata, C. elatior, C. fel-
nouilliana, C. fluminea, C. japonica,
C. orientalis, C. papyracea, C. sui-
fuensis, C. vinca: S2:113-124

Kuwait
Acanthochitona woodwardi, Chiton
peregrinus, C. (Rhyssoplax) affinis,
Falaika Island, Ischnochiton winck-
worthi, |. yerbury, Notoplax
(Notoplax) arabica, Tonicia (Lucilina)
sueziensis: 6(1):115-130

Kyles Ford, TN
Cumberlandia monodonta: 4(1):13-19

Laguna Madre, TX
Fossils, Molluscan Communities:
1:89

LaGrue Bayou, AR
Corbicula fluminea: S2:7-39

Lake Albert, Africa
Biomphalaria choanomphala, B.
smithii, B. stanleyi, B. sudanica,
Schistosoma mansoni: 5(1):85-90

Lake Allatoona, GA
Corbicula fluminea: S2:7-39

Lake Arlington, TX
Corbicula fluminea: 3(2):267-268;
$2:7-39, 99-111, 231-239. Physella
virgata virgata: S2:7-39. Quadrula
quadrula: S2:99-111 (passim)

Lake Benbrook, TX
Corbicula fluminea, Lepomis
microlophus, Minytrema melanops:
$2:7-39

Lake Buena Vista, FL
Corbicula fluminea: S2:7-39

Lake Casitas, CA
Corbicula fluminea: S2:7-39

Lake Constance (Austria, Germany,

Switzerland)

Ancylus fluviatilis: 3(2):151-168

Lake Contos, MI
Anodonta imbecilis: 4(2):231-232

Lake Edward, Africa
Biomphalaria choanomphala, B.
smithii, B. stanleyi, B. sudanica,
Schistosoma mansoni: 5(1):85-90

Lake Eorom, Denmark

Pisidium subtruncatum: 5(1):41-48
Lake Erie, MI, OH, PA, Ont.

Corbicula: $2:1-5, 125-132, 185
Lake Fairfield, TX

Corbicula: $2:125-132
Lake Hippochee, FL

Corbicula fluminea: S2:7-39
Lake Hwama, PRC

Corbicula fluminea: S2:113-124
Lake Inks, TX

Corbicula fluminea: S2:7-39
Lake Jackson, FL

Corbicula fluminea: S2:7-39
Lake Jennings, CA

Corbicula fluminea, Ictalurus fur-

catus: S2:7-39
Lake Keowee, SC

Corbicula fluminea: S2:7-39
Lake Long, TX

Corbicula fluminea: S2:179-184
Lake Lucy, FL

Corbicula fluminea: S2:7-39
Lake Lyndon B. Johnson, TX

Corbicula fluminea: S2:7-39
Lake Malawi

Bellamya jeffreysi: 4(1):107
Lake Martinez, AZ

Corbicula fluminea: S2:7-39
Lake Meade, NV

Corbicula fluminea: S2:7-39
Lake Murray, CA

Corbicula fluminea: S2:7-39
Lake Norman, NC

Corbicula fluminea: 1:96
Lake of the Pines, TX

Corbicula: $2:125-132
Lake Okeechobee, FL

Corbicula fluminea: S2:7-39
Lake Oklawaha, FL

Corbicula fluminea: S2:7-39
Lake Overholser, OK

Corbicula fluminea: S2:7-39

Anodonta piscinalis, Pisidium am-

nicum: 5(1):41-48. P casertanum, P.

conventus: 5(1):21-30
Lake Palatlakaha, FL

Corbicula fluminea: S2:7-39
Lake Piru, CA

Corbicula fluminea: S2:7-39
Lake Raven, TX

Corbicula fluminea: S2:179-184
Lake Springfield, IL

Corbicula fluminea: S2:7-39
Lake Talquin, FL

Anodonta imbecilis: 4(1):117.

Campeloma parthenum: 3(1):99. Cor-

bicula fluminea: S2:7-39. Elliptio

icterina: 1:95; 4(1):117. Villosa villosa:

1:95; 4(1):117
Lake Tanganyika, Africa

Neothauma tanganyicense, Pliodon

spekii: 4(1):107

Lake Texoma, OK, TX
Corbicula fluminea: S2:7-39
Lake Theo, TX
Anodonta grandis: S2:179-184.
Gastropoda, Unspecified: 1:99. Rana
catesbeiana: S2:179-184
Lake Thunderbird, OK
Corbicula fluminea: S2:7-39
Lake Travis, TX
Corbicula fluminea: S2:7-39
Lake Tsala, FL
Corbicula fluminea: S2:7-39
Lake Varaslampi, Finland
Sphaerium corneum: 5(1):41-48
Lake Victoria, Africa
Bellamya, B. capillata, B. jeffreysi,
Bellawya unicolor: 4(1):107. Biom-
phalaria choanomphala, B. smithii,
B. stanleyi, B. sudanica: 5(1):85-90.
Caelatura, Neothauma tanganyi-
cense, Pliodon, P. ovata, P spekii: .
4(1):107. Schistosoma mansoni:
5(1):85-90
Lake Waccamaw, NC
Corbicula: S2:125-132. C. fluminea:
3(1):100; S2:7-29, 219-222. Elliptio
cistelliformis, E. fisheriana, E.
folliculata, E. lanceolata, E. producta,
E. ravenelli, E. waccamawensis,
Lampsilis crocata, Leptodea
ochracea, Najas guadalupensis:
1:61-68. Nuphar luteum: 3(1):100.
Nuphar luteum sagittifolium, Panicum
hemitomon, Plant-Bivalve Associa-
tions, Plectonema sp., Toxolasma
pullus, Villosa ogeecheensis: 1:61-68
Lake Wylie, NC
Corbicula fluminea: S2:7-39
LAnguille River, AR
Corbicula fluminea: S2:7-39
Laos
Corbicula crocea: S2:113-124
Leaf River, MS
Corbicula fluminea: S2:7-39
Lesser Antilles
Camaenidae, Paleontology,
Pleurodonte: 3(1):102-103
Lewisville Lake, TX
Corbicula fluminea: S2:179-184
Lick River, TN
Corbicula fluminea: S2:7-39
Licking River, KY
Corbicula fluminea: S2:7-39
Licking River, OH
Corbicula fluminea: S2:7-39
Limestone Creek, AL
Corbicula fluminea: S2:7-39
Little Black River, MO
Corbicula fluminea: S2:7-39
Little Brazos River, TX
Corbicula fluminea: S2:7-39
Little Brosna River, Republic of
Ireland
Ancylus fluviatilis: 5(1):105-124

270 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Little Cypress Creek, AL
Corbicula fluminea: S2:7-39

Little Duck River, TN
Corbicula fluminea: S2:7-39

Little Grant River, WI
Lampsilis ventricosa, Lasmigona
costata: 5(2):165-171

Little Hickory Creek, TN
Lithasia pinguis: 1:27

Little Muskingum River, OH
Corbicula fluminea: S2:7-39

Little Ocmulgee River, GA
Corbicula fluminea: S2:7-39

Little Pee Dee River, SC
Corbicula fluminea: S2:7-39

Little Pigeon River, TN
Anodonta grandis, Epioblasma cap-
saeformis, Fusconaia barnesiana,
Lampsilis fasciola, L. ovata,
Lasmigona costata, Pleurobema
oviforme, Toxolasmus lividus, Villosa
iris, V. vanuxemensis: 6(2):165-178

Little Pigeon River, West Prong TN
Elliptio crassidens, E. dilatata, Epio-
blasma capsaeformis, Fusconaia
barnesiana, Lampsilis fasciola, L.
ovata, Lasmigona costata, Leptodea
fragilis, Medionidus conradicus,
Pleurobema oviforme, Potamilus
alatus, Quadrula pustulosa, Tox-
olasma lividus, Villosa iris, V. vanux-
emensis: 6(2):165-178

Little River, AR, OK
Corbicula fluminea: S2:7-39

Little River, KY
Corbicula: S2:7-39, 125-132

Little River, NC
Corbicula fluminea: S2:7-39

Little River, TN
Actinonaias pectorosa, Alasmidonta
viridus, Amblema plicata, Cumber-
landia mondonta, Elliptio dilatata,
Epioblasma capsaeformis, E. hay-
siana, E. triquetra, Fusconaia barne-
siana, F. barnesiana bigbyensis, F.
cuneolus appressa, Lampsilis car-
dium, L. fasciola, Lasmigona
costata, L. holstonia, Medionidus
conradicus, Pleurobema oviforme,
Toxolasma lividus glans: 6(1):19-37.
Unionids, Unspecified: 1:93-94.
Villosa iris, V. vanuxemensis:
6(1):19-37

Little River Canal, MO
Corbicula fluminea: S2:7-39

Little Sippewisset Marsha, MA
Melampus bidentatus: 4(1):121-122

Little South Fork River, KY
Villosa trabalis: 1:28

Little Tennessee River, TN
Actinonaias ligamentina gibba, Alas-

midonta marginata, Amblema plicata,

Anodonta grandis: 6(1):19-37. Cor-
bicula fluminea: S2:7-39. Cyclonaias

tuberculata, Cyprogenia stegaria,
Dromus dromas, Elliptio crassidens,
E. dilatata, Epioblasma arcaeformis,
E. brevidens, E. capsaeformis, E.
florentina, E. haysiana, E. propinqua,
E. stewartsoni, E. torulosa,
Fusconaia barnesiana, F. barnesiana
bigbyensis, F. barnesiana tumescens,
F. subrotunda, Hemistena lata,
Lampsilis abrupta, L. fasciola, L.
ovata, Lemiox rimosus, Leptodea
fragilis, Lexintonia dolabelloides,
Ligumia recta, Medionidus conradicus,
Obovaria retusa, O. subrotunda,
Plethobasus cooperianus, P.
cyphyus, Pleurobema coccineum, P.
cordatum, P oviforme, P. oviforme
holstonse, P plenum, P. rubrum,
Potamilus alatus, P ohioensis,
Ptychobranchus fasciolare, P subten-
tum, Quadrula cylindrica, Q.
metanevra, Q. pustulosa, Q. sparsa,
Strophitus undulatus, Villosa iris, V.
vanuxemensis: 6(1):19-37
Little Uchee Creek, AL
Corbicula fluminea: S2:7-39
Livermore Canal, CA
Corbicula fluminea: S2:7-39
Llano Grande Lake, TX
Corbicula fluminea: S2:179-184
Llano River, TX
Corbicula fluminea: S2:7-39, 179-184,
193-201
Locust Creek, PA
Margaritifera margaritifera: 4(1):13-19
Locust Fork, AL
Corbicula fluminea: S2:7-39
Logan Creek, MO
Corbicula fluminea: S2:7-39
Long Island Sound, CT, NY
Crassostrea virginica: S3:25-29
Long Key, FL
Acanthochitona andersoni, Acantho-
chiton zebra: 6(1):79-114. Costasiella
ocellifera, Elysia, E. papillosa, E.
subornata, E. tuca, Ercolania
coerulea, Tridachia crispata:
5(2):259-280
Long Key Reef, FL
Acanthochiles (Notoplax) hemphili,
Acanthochitona zebra, Cryptoconch-
us floridanus: 6(1):79-114
Long Island, Bahamas
Acanthochitona zebra: 6(1):79-114
Long Mountain Island Lake, NC
Corbicula fluminea: S2:7-39
Loosahatchie River, TN
Amblema plicata, Elliptio crassidens,
Lampsilis teres teres, Potamilus pur-
purata, Quadrula pustulosa, Q.
quaoarula, Tritogonia verrucosa:
6(1):19-37
Loudon Reservoir, TN
Corbicula fluminea: S2:7-39

Lough Inch, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124

Louisiana (LA)
Bay Champagne: 6(2):189-197. Bayou
Cocodrie, Bayou Magasille, Bayou
Sorrell: S2:7-39. Calcasieu River:
2:86; S2:7-39. Corbicula sp.: 2:86.
Mississippi River, Pearl River: S2:7-39.
Pleurocera acuta: 3(1):100. Red
River: S2:7-39. Sphaerium spp.:
2:86. Tensas River: S2:7-39. Thais
haemastoma canaliculata: 6(2):189-197.
Unionids, unspecified: 2:86

Louisiana Slope
Amygdalum politum, Calyptogena
ponderosa, Lucina atlantis, Lucinoma
atlantis, L. filosa, Pseudomiltha,
Solemya (Acharax) caribbaea,
Vesicomya caudata: $1:23-34

Lower Matecumbe Key, FL
Acanthochitona pygmaea: 6(1):79-114.
Laurencia obtusa, L. poitei, Tricolia
affinis: 4(2):185-199

Madagascar
Acanthochitona limbata: 6(1):115-130.
Cerithidea decollata: 2:1-20. Chiton
(Chiton) fosteri, C. huluensis,
Ischnochiton rufopunctatus, Notoplax
elegans: 6(1):115-130. Pupa suturalis:
5(2):243-258

Madison-Mariana Diversion Canal, AR
Corbicula fluminea: S2:7-39

Magdalena Plain
Paleontology: 2:84-85

Magnetic Island, Australia
Avicennia, Littorina filosa, L. scabra,
Metopograpsus, Rhizophora,
Thalamita crenata: 4(1):112

Magueyes Island, PR
Acanthochiles (Notoplax) hemphilli,
Acanthochitona lineata: 6(1):79-114

Main Canal, FL
Corbicula fluminea: S2:7-39

Maine (ME)
Acochlidacea: 2:95. Aeolidia
papillosa: 5(2):287-292. Cadlina
laevis: 4(2):205-216. Carcinus
maenas: 4(1):108. Catriona gymnota,
Coryphella gracilis, C. nobilis, C.
pellucida, C. salmonacea, C. verrilli,
C. verrucosa: 5(2):287-292. Crepidula
convexa, C. fornicata: 4(2):173-183.
Cuthona concinna: 5(2):287-292.
Dendronotus frondosus: 4(2):205-216.
Eubranchus tricolor, Facelina boston-
iensis: 5(2):287-292. Jeffrey’s Basin,
Jericho Bay: 6(1):1-8. Gulf of Maine:
5(2):287-292; 6(1):1-8. Littorina ob-
tusata: 4(1):108. Metridium senile:
5(2):287-292. Pseudovermis: 2:95.
Placopecten magellanicus, Ringtown
Island: 6(1):1-8. Setoaeolis pilata:
5(2):287-292. Tonicella rubra:
6(1):69-78

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 271

Malaysia
Cerithidea cingulata, C. obtusa:
2:1-20. Corbicula javanica, C. malac-
censis: S2:113-124. Ischnochiton
(Ischnochiton) winckworthi:
6(1):115-130. Perna viridis:
5(2):159-164. Tricula: 2:88

Maldive Islands
Chiton huluensis: 6(1):115-130

Manice Bayou, AR
Corbicula fluminea: S2:7-39

Manitoba, Canada
Macoma balthica: 1:90

Manora Island, Pakistan
Ischnochiton (Ischnochiton) yerburyi:
6(1):115-130

Marshall Islands
Akera soluta, Bornella anguilla,
Chromodoris geometrica, Elysia
livida, E. vatae: 5(2):243-258.
Enewetak: 5(2):197-214, 243-258.
Flabellina, Halgerda wasinensis,
Marianina rosea, Platydoris cruenta,
P. scabra: 5(2):243-258. Pleurehdera
haraldi: 5(2):197-214

Marthas Vineyard, MA
Crepidula convexa, C. fornicata, C.
plana, Limulus polyphemus, Littorina
littorea, Lunatia heros: 3(1):33-40

Maryland (MD)
Broad Creek: S3:25-29. Chesapeake
Bay: S2:7-39. Choptank River, Cor-
bicula fluminea: S2:7-39. Crassostrea
virginica: $3:25-29. Elliptio
fisherianus, E. lanceolata, E. produc-
tus: 3(1):94. Nassawango Creek,

Potomac River: S2:7-39. Spisula con-

fraga: 4(1):39-42. Susquehanna
River: S2:7-39. Teinostoma nana:
4(1):39-42. Tred Avon River:
$3:25-29. Wicomico River: S2:7-39

Mashta Island, FL
Alvania auberiana, Caecum nitidum,
Granulina ovuliformis, Halodule
wrightii, Laurencia poitei, Marginella
aureocincta, Rissoella caribaea, Ris-
soina bryerea, Smaragdia viridis
viridemaris, Thalassia testudinum,
Tricolia thalassicola, Zebina browni-
ana: 4(2):185-199

Masirah Island, Oman
Callistochiton adenensis, Chiton
(Chiton) peregrinus: 6(1):115-130

Massachusetts (MA)
Aeolidia papillosa: 4(2):205-216. Am-
nicola limosa: 5(1):9-19. Arctica
islandica: $3:51-57. Arenicola: 2:96.
Buzzards Bay; 6(1):69-78.
Campeloma decisum: 5(1):9-19.
Chaetopleura apiculata: 6(1):69-78.
Cipangopaludina chinensis: 5(1):9-19.

Coryphella salmonacea: 4(2):205-216.

Crepidula convexa: 3(1):33-40;
4(2):173-183. C. fornicata: 3(1):33-40.

C. plana: 3(1):33-40; 4(2):173-183.
Cuttyhunk Island: S3:51-57. Ensis:
2:96. Ferrissia fragilis, F. parallela:
5(1):9-19. Gemma gemma, Glycera:
2:96. Gyraulus circumstriatus, G.
deflectus, G. parvus, Helisoma
anceps, H. campanulatum, H.
trivolvis, Laevapex fuscus: 5(1):9-19.
Leptosynapta: 2:96. Limulus poly-
phemus: 2:96; 3(1):33-40. Little Sip-
pewisset Marsh: 4(1):121-122;
4(2):236. Littorina littorea, Lunacia
heros: 3(1):33-40. Lyrogyrus granum,
L. pupoidea: 5(1):9-19. Margaritifera
margaritifera: 4(1):13-19. Marthas
Vineyard: 3(1):33-44. Melampus
bidentatus: 4(1):121-122; 4(2):236.
Mercenaria: 2:96. Micromentus
dilatatus: 5(1):9-19. Mya, Nereis: 2:96.
Nucella lapillus: 4(2):201-203. Physa
ancillaria, P_ heterostropha, Planor-
bula armigera, Promenetus ex-
acuous, Pseudosuccinea columella:
5(1):9-19. Scoloplos, Solemya velum:
2:96. Stagnicola elodes: 5(1):9-19.
Syllis: 2:96. Tonicella rubra:
6(1):69-78. Valvata tricarinata,
Viviparius georgianus: 5(1):9-19.
Woods Hole: 3(1):33-40; 6(1):69-78

Matagorda Bay, TX
Periploma margaritaceum, P.
orbiculare: 2:35-40

Maui, HI
Barleeia: 4(2):232-233

Maumee River, OH
Corbicula fluminea: S2:7-39, 185

Mauritius
Bulinus cernicus: 1:107. Elysia
moebii, E. virgata, Halgerda formosa,
Mypselodoris carnea: 5(2):243-258.
Onithochtion maillardi: 6(1):115-130.
Pleurobranchus inhacae:
5(2):243-258. Shistosoma
haematobium: 1:107

Mayakka River, FL
Corbicula fluminea: S2:7-39

Mayberry Cut, CA
Corbicula fluminea: S2:7-39

McKinney Bayou, AR
Corbicula fluminea: S2:7-39

McMahan Site, TN
Actinonaias ligamentina, Alasmidonta
marginata, A. viridis, Amblema
plicata, Anodonta grandis,
Campeloma decisum, Cyclonaias
tuberculata, Cyprogenia stegaria,
Dallas Component, Dromus dromas,
Elliptio crassidens, E. dilatata,
Epioblasma arcaeformis, E.
brevidens, E. capsaeformis, E. floren-
tina, E. haysiana, E. stewardsoni, E.
torulosa, Fusconaia subrotunda,
Hemistena lata, lo fluvialis, Lampsilis
fasciola, L. ovata, Lasmigona costata,

L. holstonia, Lemiox rimosus, Lep-
toxis praerosa, Lexingtonia
dolabelloides, Ligumia recta, Lithasia
(Angitrema) verrucosa, Medionidus
conradicus, Obovaria subrotunda,
Plethobasus cooperianus, P.
cyphyus, Pleurobema cordatum, P
oviforme, P. plenum, P. rubrum,
Pleurocera canaliculatum, P parvum,
Potamilus alatus, Ptychobranchus
fasciolaris, P subtentum, Quadrula
cylindrica, Q. pustulosa, Q. sparsa,
Toxolasma lividus, Villosa iris, V.
trabalis: 6(2):165-178

Media Luna Reef
Acanthochitona lineata, A. pygmaea:
6(1):79-114

Mediterranean Sea
Atagema gibba, A. rugosa, Berthella
plumula, Chelidoneura hirundinina:
5(2):243-258. Chiton huluensis, C.
(Rhyssoplax) olivaceus: 6(1):115-130.
Chromodoris krohni: 5(2):185-196.
Doriopsilla miniata, Doto coronata, D.
pinnatifida, D. rosea, Elysia viridus,
Goniodoris castanea: 5(2):243-258.
Hypselodoris bilineata, H. gracilis, H.
messinensis, H. valenciennesi:
5(2):185-196. Kalopocamus ramosus,
Limacia clavigera, Lobiger souver-
biel: 5(2):243-258. Mexichromis
tricolor: 5(2):185-196. Octopus
vulgaris: 6(1):45-48. Patella perversa:
5(2):197-214. Placida dendritica:
5(2):243-258. Pleurobranchaea
meckelii: 5(2):197-214. Polycera quad-
rilineata, Prutfolia pselliotes:
5(2):243-258. Scaeurgus unicirrhus:
6(2):207-211. Tergipes tergipes:
5(2):243-258. Theba pisana: 1:104.
Thecacera pennigera, Umbraculum
Sinicum: 5(2):243-258

Meigs Creek, OH
Corbicula fluminea: S2:7-39

Meramec River, MO
Corbicula fluminea: S2:7-39.
Cumberlandia monodonta: 4(1):13-19

Merced River, CA
Corbicula fluminea: S2:7-39

Mexico
Acanthochitona andersoni, A.
pygmaea: 6(1):79-114. Amiantus sp.:
4(1):1-12. Ammonitellidae: 1:97.
Andara (Esmerarca) sp., A. nux:
4(1):1-12. Arcticacea: 3(1):103. Baja
California: 1:97; 3(1):102-103. Baja
California del Norte: 3(1):103. Baja
California Sur: 3(1):103; 4(1):1-12.
Bernardina, B. bakeri, B. margarita,
Bernardinidae: 3(1):103. Bulimulidae:
1:97; 4(1):113-114. Calliostoma han-
nibali, Calyptraea sp.: 4(1):1-12.
Campeche: 2:1-20; 6(1):79-114. Car-
dita (Cardites) sp.: 4(1):1-12.

272

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Cerithidea montagnei, C. pliculosa:
2:1-20. Cerithium sp., Chione
(Chione) richthofeni, C. (Chionopsis)
sp., C. sp., Choromytilus palliopunc-
tatus: 4(1):1-12. Choya Bay:
6(1):45-48. Chromodoris annulata:
5(2):243-258. Crassilabrum wittichi,
Crassispira starri: 4(1):1-12. Crasso-
strea cortiezensis, C. rhizophorae, C.
virginica: 1:108. Crepidula sp.,
Crucibulum scutellatum: 4(1):1-12.
Cyamiacea: 3(1):103. Cycinellas sp.,
Cymia heimi, Cypraea amandus,
Divalinga comis, Drillia (Clathrodrilia)
sp.: 4(1):1-12. Halodakra, H.
salmonea, H. (Halodakra) subtrigona:
3(1):103. Haplotrematidae: 1:97.
Helminthoglypta ayersiana, H.
(Charodotes) traskil: 3(1):103. Helmin-
thoglyptidae: 1:97; 3(1):102-103. Hex-
aplex erythrostomus: 6(1):45-48. Hip-
ponix pilosus: 4(1):1-12. Holocene:
4(2):238-239. Isla Mujere: 6(1):79-114.
Jalisco: 2:1-20; 3(1):103. Knefastia
sp., Lucina (Lucinisca) sp.: 4(1):1-12.
Lysinoe, L. ghiesbreghti: 3(1):102-103.
Macron hartmani, Melongena
melongena, M. melongena consors:
4(1):1-12. Muricanthus nigritus:
6(1):45-48. Mytilus canoasensis
vidali, Nassarius versicolor, Neverita
(Glossaulax) andersoni: 4(1):1-12.
Nucella emarginata: 1:105. Nuevo
Leon: 3(1):102-103. Octopus digueti:
6(1):45-48. Oreohelicidae: 1:97.
Orymaeus: 4(1):113-114. Ostrea sp.:
4(1):1-12. Paleontology: 2:84-85;
3(1):98, 102-103; 4(1):1-12;
4(2):238-239. Penninsula Effect: 1:97.
Plicatula inezana: 4(1):1-12. Pliocene:
4(2):238-239. Protothaca sp.:
4(1):1-12. Punta Palmar, Quintana
Roo: 6(1):79-114. Rhabaotus, R.
baileyi, R. nigromontanus:
4(1):113-114. Raeta sp., San Ignacio
Formation, Sanguinolaria toulai,
Siphocypraea henekeni, Siphonaria
maura pica, Solenosteira sp.:
4(1):1-12. Sonora: 4(1):113-114;
6(1):45-48. Speciation: 1:97. Spiraci-
dae: 1:9. Strombina sp., Tegula sp.,
Terebra burckhardti: 4(1):1-12. Thais
emarginata: 1:105. Theodoxus sp.,
Trachycardium sp., Trochita radians,
T. spirata, T. trochiformis: 4(1):1-12.
Turridae: 3(1):98. Turritella abrupta, T.
altilira, T. bifastigata, T. bosei, T.
costaricensis, T. crocus, T. inezana
bicarina, Vermetus contortus:
4(1):1-12. Xerarionta: 3(1):102-103.
Yucatan Peninsula: 1:108;
6(1):79-114. Yucum Balam: 6(1):79-114

Miami River, OH

Corbicula fluminea: S2:7-39

Michigan (Ml)
Actiononaias carintata: 3(1):105. A.
ellipsiformis: 3(1):93. Alasmidonta
marginata, A. viridis, Amblema
plicata: 3(1):105. Anodonta grandis:
3(1):93. A. grandis grandis: 3(1):105.
A. imbecilis: 3(1):93, 105;
4(2):231-232. Anodontoides ferussac-
janus: 3(1):93, 105. Caruncula parva:
3(1):105. Cedar River: 3(1):93;
4(2):231-232. Corbicula: S2:1-5. C.
fluminea: S2:7-39. Cyclonaias tuber-
culata, Detroit River, Dysnomia
sulcata delicata, D. torulosa
rangiana, D. triquetra, Elliptio dilatata:
3(1):105. Fusconaia flava: 3(1):93,
105. F subrotunda: 3(1):105.
Helisoma anceps: 5(1):73-84. Lake
Contos: 4(2):231-232. Lake Erie:
$2:1-5, 7-39. Lampsilis fasciola:
3(1):105. L. ovata, L. radiata: 3(1):93.
L. radiata luteola, L. ventricosa,
Lasmigona complanata: 3(1):105. L.
compressa 3(1):93, 105. L. costata,
Leptodea fragilis, L. leptodon,
Ligumia nasuta, L. recta: 3(1):105.
Limnaea (Stagnicola) elodes: 1:67-70.
Obliquaria reflexa, O. olivaria, O.
subrotunda;: 3(1):105. Physa integra:
5(1):73-84. Pleurobema coccineum,
Proptera alata, Ptychobranchus
fasciolare, Quadrula pustulosa, Q.
quadrula: 3(1):105. Sandy Creek:
$2:7-39. Simpsoniconcha ambigua,
Strophitus undulatus, Truncilla
donaciformis; T. truncata, Villosa
fabalis, V. iris: 3(1):105.

Mid-Atlantic Bight
Illex illecebrosus, Loligo peali:
4(1):101

Middle River Canal, FL
Corbicula fluminea: S2:7-39

Midway Island
Alvania (Alvania) isolata, Barleeia,
Euplica turturina: 4(2):232-233

Milford Haven, VA
Crassostrea virginica, Haplosporidium
nelsoni: S3:17-23

Millville Site, WI
Actinonaias ligamentina carinata,
Amblema plicata, Elliptio dilatata,
Fusconaia ebena, F. flava, Pletho-
basus cyphus, Quadrula metanerva:
5(2):165-171

Minnesota (MN)
Actinonaias ligamentina carinata,
Alasmidonta marginata, Amblema
plicata plicata, Anodonta grandis
corpulenta, A. grandis grandis, A.
imbecilis, A. suborbiculata, Arcidens
confragosus: 1:51-60. Corbicula:
$2:1-5. C. fluminea: S2:7-39.
Cyclonaias tuberculata, Ellipsaria
lineolata, Elliptio crassidens crassidens,

E. dilatata, Fusconaia ebena, F.
flava, Hendersonia occulta, Lampsilis
higginsi, L. radiata luteola, L. teres
anodontoides, L. teres teres, L.
ventricosa, Lasmigona complanata,
L. costata, Leptodea fragilis, Ligumia
recta, Magnonaias nervosa: 1:51-60.
Minnesota River: S2:7-39. Mississip-
pi River, Obovaria olivaria, Pletho-
basus cyphyus, Pleurobema rubrum,
P. sintoxia, Potamilus alatus, P. ohien-
sis, Quadrula metanerva, Q.
nodulata, Q. pustulosa pustulosa, Q.
quaarula: 1:51-60. St. Croix River:
$2:1-5. Strophitus undulatus un-
dulatus, Toxolasma parvus, Tritogonia
verrucosa, Truncilla donaciformis, T.
truncata: 1:51-60

Minnesota River, MN

Corbicula fluminea: S2:7-39

Mississippi (MS)

Allan Branch, Amite River: S2:7-39.
Anodonta imbecilis: 4(1):21-23, Bear
Creek, Big Black Creek, Big Black
River, Bouge Phalia River, Buckatun-
na Creek, Buttahatchie River, Chick-
asawhay River, Chunky River, Cold-
water River: S2:7-39. Corbicula
fluminea: 2:87; 4(1):21-23; 4(2):234;
S2:7-39. Elliptio crassidens,
Fusconaia flava: 4(1):21-23. Geuken-
sia demissa granosissima: 4(1):112;
5(2):173-176. Halstead Bayou:
6(2):199-206. Lampsilis claibornensis,
L. ovata ventricosa, L. radiata
luteola, L. straminea claibornensis,
L. teres anodontoides: 4(1):21-23.
Leaf River: S2:7-39. Leptodea
fragilis: 4(1):21-23. Mississippi River,
Moss Creek: S2:7-39. Obovaria
subrotunda: 4(1):21-23. Old Fort
Bayou: 6(2):199-206. Okatibee
Creek, Okatoma Creek, Pascagoula
River, Pearl River: S2:7-39. Poly-
mesoda caroliniana: 6(2):199-206. P
carolinianus: 4(2):234. Potamilus pur-
purata, Quadrula pustulosa:
4(1):21-23. Shubuta Creek,
Souinlovey Creek, Steel Bayou:
$2:7-39. Strophitus subvexus:
4(1):21-23. Sunflower River: S2:7-39.
Tallahalla Creek: 2:87; S2:7-39. Tib-
bee Creek, Tombigbee River:
S2:7-39. Toxolasma texasensis,
Tritogonia verrucosa, Uniomerus
declivus, Villosa lienosa: 4(1):21-23.
Woodward Creek, Yalobusha River,
Yazoo River, Yockanookany River:
$2:7-39

Mississippi River

Actinonaias ligamentina carinata,
Alasmidonta marginata: 1:51-60.
Amblema plicata plicata: 1:51-60;
6(1):49-54. Anodonta grandis

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 273

corpulenta, A. grandis grandis, A.
imbecilis: 1:51-60. A. suborbiculata:
1:51-60; 4(2):230-231. Arcidens con-
fragosus: 1:51-60; 5(2):165-171. Cor-
bicula: S2:1-5. C. fluminea: S2:7-39.
Cumberlandia monodonta: 4(1):13-19.
Cyclonaias tuberculata, Ellipsaria
lineolata, Elliptio crassidens
crassidens, E. dilatata: 1:51-60.
Fusconaia ebena: 1:51-60;
5(2):165-171. Fusconaia flava,
Hendersonia occulta: 1:51-60. Illinois,

lowa: S2:1-5, 7-39. Lampsilis higginsi:

1:51-60; 4(2):230; 6(1):39-43, 49-54.
L. radiata luteola: 1:51-60;
4(2):230-231. L. teres anodontoides:
1:51-60; 5(2):165-171. L. teres teres,
L. ventricosa, Lasmigona com-
planata, L. costata, Leptodea fragilis,
Ligumia recta: 1:51-60. Magnonaias
nervosa: 1:51-60; 4(2):230-231. Min-
nesota: 1:51-60; 4(2):230-231.
Missouri: 4(1):13-19. Obovaria
Olivaria: 1:51-60; 4(2):230-231.
Plethobasus cooperianus: 6(1):49-54.
P. cyphyus, Pleurobema rubrum, P
sintoxia, Potamilus alatus: 1:51-60. P
capax: 4(2):230-231. P ohiensis:
1:51-60. Quaarula fragosa:
4(2):230-231. Q. metanerva: 1:51-60;
4(2):230-231. Q. nodulata, Q.
pustulosa pustulosa, Q. quadrula,
Strophitus undulatus undulatus, Tox-
olasma parvus, Tritogonia verrucosa,
Truncilla donaciformis, T. truncata:
1:51-60. Wisconsin: 1:51-60; 4(2):230,
230-231; 5(2):165-171

Missouri (MO)
Allogona profunda, Anguispira alter-
nata, A. kochi: 1:97-98. Big Creek,
Big River, Black River, Bourbeuse
River, Bryant Creek, Cane Creek:
$2:7-39. Cepaea hortensis, C.
nemoralis: 1:97-98. Corbicula
fluminea: S2:7-39. Cumberlandia
mondonta: 4(1):13-19. Current River,
Cyclonaias tuberculata, Fusconaia
ozarkensis: 2:85. Gasconade River:
$2:7-39. Helix aspersa, H. pomacea:
1:97-98. Hendersonia occulta: 1:99.
Jacks Ford River, Lampsilis
orbiculata, L. reeviana: 2:85. Little
Black River, Little River Canal,
Logan Creek, Meramec River:
S2:7-39. Mesodon clausus, M.
elevatus, M. thyroidus: 1:97-98.
Mississippi River, Missouri River,
$2:7-39. Mollusca, unspecified:
4(1):119. Moreau River, Osage River:
$2:7-39. Ozark Mountains: 4(1):119.
Pleurobema coccineum, Ptycho-
branchus occidentalis: 2:85. St.
Francis River: S2:7-39. Succinea
ovalis: Thomas Hill Reservoir:

$2:7-39. Triodopsis albolabris alleni,

T. multilineata: 1:97-98. Villosa iris

iris: 2:85. Whitewater River: S2:7-39
Missouri Key, FL

Acanthochitona andersoni, Crypto-

conchus floridanus: 6(1):79-114
Missouri River

Anodonta grandis corpulenta, A.

grandis grandis, A. suborbiculata:

1:71-74. Corbicula fluminea: S2:7-39.

Lampsilis teres teres, Lasmigona

complanata, Leptodea fragilis, L. lep-

todon, Potamilus alatus, P. ohiensis,
Quadrula quadrula, Tritogonia ver-
rucosa, Truncilla donaciformis, T.
truncata: 1:71-74
Mobile River, AL
Corbicula fluminea: S2:7-39
Mobile River System
Lampsilis altilis, L. perovalis: 1:94
Mobjack Bay, VA

Crassostrea virginica, Haplosporidium

nelsoni: S3:17-23
Mohave Desert

Cooper, James Graham: 1:89
Mokelumne Aqueduct, CA

Corbicula fluminea: S2:7-39
Mokelumne River, CA

Corbicula fluminea: S2:7-39
Moluccas

Chiton huluensis: 6(1):115-130
Monongahela River, WV

Corbicula fluminea: S2:7-39
Monte Alto Reservoir, TX

Corbicula fluminea: S2:7-39
Moorea Island, French Polynesia

Partula mooreana, P. suturalis:

1:103-104
Moreau River, MO

Corbicula fluminea: S2:7-39
Mosquitoe Creek, FI

Corbicula fluminea: S2:7-39. Anodon-

ta imbecilis: 4(2):231-232
Moss Creek, MS

Corbicula fluminea: S2:7-39
Mozambique

Cancellaria lamellosa: 2:57-61.

Chiton (Chiton) fosteri, C. huluensis,

Ischnochiton kilburni, |. sansibaren-

sis, |. (Ischnochiton) yerburyi, I.

(Radsiella) delagoaensis,

Onithochiton litteratus, Tonicia

(Lucilina) carnosa: 6(1):115-130
Mud Creek, AL

Corbicula fluminea: S2:7-39
Mud River, KY

Corbicula fluminea: S2:7-39
Mud River, WV

Corbicula fluminea: S2:7-39
Murder Creek, AL

Corbicula fluminea: S2:7-39
Muskingum River, OH

Corbicula fluminea: S2:7-39.

Unionids, Unspecified: 1:93

Nanticoke River, DE
Corbicula fluminea: S2:7-39

Napier Range, Australia
Amplirhagada: 1:98-99.
Westraltrachia: 1:98-99

Nassawango Creek, MD
Corbicula fluminea: S2:7-39

Natal
Onithochiton litteratus: 6(1):115-130

Nebraska (NB)
Anodonta grandis corpulenta, A.
grandis grandis, A. suborbiculata:
1:71-74. Cionella lubrica: 3(1):27-32.
Lasmigona complanata, Leptodea
fragilis: 1:71-74. Physella varigata
varigata: 3(2):243-265. Potamilus
ohiensis, Quadrula quadrula,
Tritogonia verrucosa, Truncilla trun-
cata: 1:71-74

Neely Henry Lake, AL
Corbicula fluminea: S2:7-39

Negev Desert
Theba pisana: 1:104

Neuse River, NC
Elliptio complanata: 3(1):104-105. E.
fisherianus, E. lanceolata, E. produc-
tus: 3(1):94

Nevada (NV)
Corbicula fluminea, Lake Meade:
$2:7-39

New Brunswick, Canada
Melampus bidentatus: 4(1):121-122;
4(2):236. Neopanope Sayi: S3:59-70

New Caledonia
Phyllodesmium piondimiei:
5(2):243-258

New England (US)
Amnicola limosa: 3(1):99. Crepidula
convexa, C. fornicata, C. plana: 1:110.
Ferrissia fragilis, Fossaria modicella,
Gyraulus circumstriatus, G. deflectus,
Helisoma anceps, H. campanulatum,
Laevapex fuscus, Micromenetus
dilatatus: 3(1):99. Onchidoris aspera:
5(2):293-301. Physella anceliaria,
Planorbula armigera, Promentetus
exacusous, Pseudosuccinea col-
umella: 3(1):99

New Guinea
Corbicula debilis: S2:113-124. Perna
viridis: 5(2):159-164

New Hebrides Islands
Paraganitus ellynnae: 5(2):281-286

New Jersey (NJ)
Acteon wetherii: 4(1):39-42. Bankia
gouldi: 4(1):89-99; S1:101-109.
Barnegat Bay: 4(1):89-99; S1:101-109.
Boveria teredinidi, B. zeukevitchi:
$1:101-109. Corbicula fluminea:
3(1):100-101; S2:1-5, 7-39.
Crassostrea: S1:101-109. Delaware
River: S2:7-39. Haplosporidium:
$1:101-109. Limnodrilus: S2:7-39.
Mulinia lateralis: 4(1):39-42. Oyster

274

New

New

New

New

New

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Creek: 4(1):89-99. Peloscolex ferox,
Procladius culiciformis: S2:7-39.
Raritan River: 3(1):100-101.
Sphaerium transversum: S2:7-39.
Teredo bartschi: 4(1):89-99;
$1:101-109. T. furcifera: S1:101-109. T.
navalis: 4(1):89-99; S1:101-109
Mexico (NM)

Cabelle Reservoir, Corbicula
fluminea, Elephant Butte Reservoir,
Pecos River, Rio Grande, West
Drain: S2:7-39

River, VA, WV

Corbicula: S2:1-5. C. fluminea:
4(1):116; S2:7-39, 69-81. Lasmigona
subviridis: 6(2):179-188

South Wales, Australia
Onithochtion quercinus, O.

rugulosus, O. scholvieni: 6(1):115-130.

Tylodina corticalis, Umbraculum um-
braculum: 5(2):197-214

York (NY)

Amnicola limosa: 5(1):9-19.
Campeloma decisum: 5(1):9-19,
101-104. Cipangopaludina chinensis:
5(1):9-19. Bithynia tentaculata:
3(2):179-186. Crepidula convexa, C.
plana: 4(2):173-183. Donax fossor:
3(1):92. Ferrissia fragilis, F. parallela:
5(1):9-19. Gastropoda, unspecified:
5(1):101-104. Gyraulus circumstriatus,
G. deflectus, G. parvus, Helisoma
anceps, H. campanulatum: 5(1):9-19.
H. trivolvis: 4(2):229; 5:9-19.
Laevapex fuscus: 5(1):9-19. Leptoxis
carinata: 3(2):169-177. Lyrogyrus
granum, L. pupoidea, Micromentus
dilatatus, Physa ancillaria, P
heterostropha, Planorbula armigera,
Promenetus exacuous, Pseudosuc-
cinea columella, Stagnicola elodes,
Valvata tricarinata: 5(1):9-19.
Viviparus georgianus: 3(2):268;
5(1):9-19

Zealand

Bathyberthella zelandiae:
5(2):197-214. Bursatella leachii:
5(2):243-258. Offadesma angasi:
2:35-40. Perna canaliculus:
5(2):159-164. Philine angasi, P
auriformis: 5(2):185-196. Polycera
hedgpethi: 5(2):243-258. Pseudo-
succinea columella: 5(1):9-19.
Pseudovermis hancocki:
5(2):281-286. Rostanga muscula,
Thecacera pennigera: 5(2):243-258

Newfoundland, Canada

Illex illecebrosus: 2:51-56

Newport River, NC

Chaetopleura apiculata, Diodora
cayenensis: 4(1):107-108

Nicaragua

Mitrridae, Pleioptygma, P. helenae,
Volutidae: 3(1):97-98

Nicobares Islands

Pleurobranchella nicobarica:
5(2):243-258

Nine Mile Creek, TN

Corbicula fluminea: S2:7-39

No Name Key, FL

Acanthochitona pgymaea: 6(1):79-114

Nolichucky River, TN

Actinonaias ligamentina, A. ligamen-
tina gibba, A. pectorosa, Alasmidon-
ta marginata, Amblema plicata:
6(1):19-37. Corbicula fluminea:
$2:7-39. Cumberlandia monodonta:
6(1):19-37. Cyclonaias tuberculata
tuberculata, Elliptio crassidens, E.
dilatata, Epioblasma capsaeformis,
E. torulosa gubernaculum, E. tri-
quetra, Fusconaia barnesiana, F.
cuneolus appressa, F. subrotundaa, F.
subrotunda lesuerianus, Lampsilis
cardium, L. fasciola, L. ovata,
Lasmigona costata, L. holstonia,
Pleurobema cordatum, Potamilus
alatus, Ptychobranchus fasciolare,
Quadrula intermedia, Q. pustulosa,
Truncilla truncata, Villosa fabalis, V.
iris, V. vanuxemensis: 6(1):19-37

Nolin River, KY

Corbicula fluminea: S2:7-39

Nore River, Republic of Ireland

Ancylus fluviatilis: 5(1):105-124

North America

Crassatella ponderosa, C. vadosa,
Cretaceous, Megapallifera, Pachy-
thaerus, Pallifera, Philomycus:
4(2):238

North American Basin

Prochaetodermatidae: 3(1):97

North Bimini Island

Acanthochitona andersoni:
6(1):79-114. Thalassia testudinum,
Tricolia bella: 4(2):185-199

North Canadian River, OK

Corbicula fluminea: S2:7-39

North Carolina (NC)

Anodonta implicata: 3(1):104-105.
Beaufort Inlet, Bogue Sound:
4(1):107-108. Cape Fear River:
S2:7-39. Cashie River: 3(1):104-105.
Catawba River: S2:7-39, 125-132.
Chaetopleura apiculata: 4(1):107-108.
Chowan River: S2:219-222. Cor-
bicula: $2:125-132. C. fluminea: 1:96;
3(1):100, 104-105; S2:7-39, 219-222.
Diodora cayenensis: 4(1):107-108.
Eden River: S2:7-39. Elliptio
angustata: 1:95. E. angustatus:
3(1):94. E. (Canthyria) steinstansana:
3(1):104-105. E. cistelliformis: 1:61-68.
E. complanata: 3(1):104-105. E. em-
monsi: 3(1):94. E. fisheriana: 1:61-68.
E. fisherianus: 3(1):94. E. foliculata:
1:61-68. E. folliculatus, E. hazelhursti-
anus: 3(1):94. E. lanceolata: 1:61-68,

1:94-95, 1:95; 3(1):94. E. producta:
1:61-68. E. productus: 3(1):94. E.
ravenelli: 1:61-68. E. shepardiana, E.
subcylindraceus: 3(1):94. E. wac-
camawensis: 1:61-68. Hendersonia
occulta: 1:99. Ischnochiton striolatus:
4(1):107-108. Lake Norman: 1:96.
Lake Waccamaw: 1:61-68; 3(1):100;
$2:7-39, 125-132, 219-222. Lampsilis
crocata: 1:61-68. Leptodea ochracea:
1:61-68; 3(1):104-105. Ligumia nasuta:
3(1):104-105. Little River: S2:7-39.
Long Mountain Island Lake: S2:7-39.
Najas guadalupensis: 1:61-68. Neuse
River: 3(1):94, 104-105. Newport
River: 4(1):107-108. Nuphar luteum:
3(1):100. N. /Juteum sagittifolium,
Panicum hemitomon, Plant-Bivalve
Associations, Plectonema sp.:
1:61-68. Richardson Creek: S2:7-39.
Roanoke River: 3(1):104-105. Rocky
River: S2:7-39. Tar River: 1:95-95,
1:95; 3(1):94, 104-105. Toxolasma
pullus: 1:61-68. Uhwarrie River:
$2:7-39. Villosa ogeecheensis:
1:61-68. Waccamaw River: S2:7-39.
Wateree River: S2:125-132

North Fork Creek, TN

Corbicula fluminea: S2:7-39

North Fork Obion River, TN

Amblema plicata, Anodonta plicata
plicata, Arcidens confragosus,
Fusconaia ebena, F. flava, F. flava
trigona, Lampsilis cardium satura, L.
teres teres, Lasmigona complanata,
Megalonaias nervosa, Plectomaris
dombeyanus, Quadrula pustulosa
mortoni, Q. quadrula, Tritogonia ver-
rucosa, Truncilla truncata: 6(1):19-37

North Mosquito Creek, FL

Corbicula fluminea: S2:7-39

North River, AL

Corbicula fluminea: S2:7-39

Norton Sound, AK

Asterias amurensis, Macoma
calcarea, Mya truncata, Serripes
groenlandicus, Yoldia hyperborea:
2:94

Norwegian Sea

Onchidoris muricata, O. varians:
2:94

Notchy Creek, TN

Corbicula fluminea: S2:7-39

Nova Scotia, Canada

Crassostrea virginica: S3:25-29

Nueces River, TX

Corbicula fluminea: S2:7-39

Nuevo Leon, Mexico

Lysinoe ghiesbreghti, Paleontology:
3(1):102-103

Obey River, TN

Actinonaias ligamentina, A. pectorosa,
Alasmidonta marginata, Amblema
plicata: 6(1):19-37. Corbicula

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 275

fluminea: S2:7-39. Cyclonaias tuber-
culata, Elliptio crassidens, E. dilatata,
Epioblasma capsaeformis, E. floren-
tina, E. florentina walkeri, E. triquetra,
Fusconaia subrotunda, Lampsilis
abrupta, L. fasciola, L. ovata, L.
teres anodontoides, Lasmigona
costata, Ligumia recta latissima,
Obliquaria reflexa, Obovaria
subrotunda, Pleurobema oviforme,
Potamilus alatus, Ptychobranchus
fasciolare, P subtentum, Quadrula
cylindrica, Q. metanerva, Strophitus
undulatus, Tritogonia verrucosa,
Villosa iris, V. taeniata: 6(1):19-37. V.
trabalis: 1:28; 6(1):19-37
Ochlocknee River, FL
Campeloma geniculum: 3(1):99. Cor-
bicula fluminea, Lampsilis
claibornensis: S2:7-39
Ocmulgee River, GA
Anodonta imbecilis: 4(2):231-232.
Corbicula fluminea, Lampsilis
anodontoides floridensis, L. uniomin-
atus, Quincucina infucata: S2:7-39
Ogeechee River, GA
Corbicula fluminea: S2:7-39
Ohio (OH)
Brush Creek: S2:7-39. Corbicula:
$2:1-5, 125-132. C. fluminea: 3(1):94;
4(1):81-88; S2:7-39, 185. Great Miami
River: 3(1):94; S2:125-132. Helisoma
anceps, H. trivolvis: 4(1):118-119.
Hocking River: S2:7-39. Lake Erie:
$2:1-5, 125-132, 185. Lasmigona
costata: 2:82. Licking River, Little
Muskingum River: S2:7-39. Maumee
River: S2:7-39, 185. Meigs Creek,
Miami River, Muskingum River, Ohio
River, Olentangy River, Olive Green
Creek, Scioto River: S2:7-39.
Sphaerium striatinum: 3(2):201-212
(passim). Stillwater River: S2:7-39.
Triodopsis tridentata tridentata: 1:98
Ohio River, IL, IN, KY, OH, PA, WV
Corbicula fluminea: 4(1):81-88;
$2:7-39. Fusconaia ebena:
5(2):177-179; 6(1):49-54
Ohoopee River, GA
Corbicula fluminea: S2:7-39
Okatibee Creek, MS
Corbicula fluminea: S2:7-39
Okatoma Creek, MS
Corbicula fluminea: S2:7-39
Okatuppa Creek, AL
Corbicula fluminea: S2:7-39
Okinawa
Cerithidea rhizophorarum: 2:1-20.
Phyllodesmium hyalinum:
5(2):243-258. Pleistocene: 2:1-20
Oklahoma (OK)
Arkansas River, Caddo Creek, Cor-
bicula fluminea, Little River, North
Canadian River, Red River: S2:7-39

Oklawaha River, FL
Corbicula fluminea: S2:7-39
Old Fort Bayou, MS

Polymesoda caroliniana: 6(2):199-206

Olentangy River, OH
Corbicula fluminea: S2:7-39
Olive Green Creek, OH
Corbicula fluminea: S2:7-39
Oman
Acanthopleura vaillantii, Al Bastan
Island, Callistochiton adenensis,
Chiton (Chiton) fosteri, C. (Chiton)
peregrinus, C. (Rhyssoplax) affinis,
Masirah Island, Onithochiton
erythraeus: 6(1):115-130
Oneida Lake, NY
Bithynia tentaculata: 3(2):179-186.
Campeloma decisum, Gastropoda,
unspecified: 5(1):101-104
Onion Creek, TX
Corbicula fluminea: S2:179-184
Ontario, Canada
Amnicola limosa, Anodonta grandis,
Campeloma decisum, Cincinnatia
cincinnatiensis, Elliptio complanata,
Gyraulus parvus, Helisoma anceps,
Lampsilis radiata, Musculium securis,
Physella gyrina, Pisidium casertanum,
P. compressum, P. ferrugineum, P
variable, Sphaerium rhomboideum,
S. simile, S. striatinum, Valvata
tricarinata: 5(1):31-39
Oostanula River, GA
Corbicula fluminea: S2:7-39
Oregon (OR)
Ancipenser transmontanus, Colum-
bia River, Corbicula fluminea:
$2:7-39. Halodakra salmonea:
3(1):103. John Day River: S2:7-39.
Loligo opalescens: 4(2):239. Smith
River, Suislaw River, Umpqua River,
Willamette River: S2:7-39
Osage River, MO
Corbicula fluminea: S2:7-39
Ouachita River, AR
Corbicula fluminea: S2:7-39
Owen Doher River, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124
Owens River, CA
Corbicula fluminea: S2:7-39
Owenwee River, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124
Oyster Creek, NJ
Teredo bartschi: 4(1):89-99
Ozark Mountains, MO
Mollusca, unspecified: 4(1):119
Paint Rock River, AL
Corbicula fluminea: S2:7-39
Pakistan
Ischnochiton haersoltei, |. (Ischno-
chiton) winckworthi, |. (Ilschnochiton)
yerburyi, Marmora Island:
6(1):115-130. Thecacera pennigera:
5(2):243-258

Palm Beach Inlet, FL
Acanthochitona andersoni, A.
balesae, A. roseojugum: 6(1):79-114

Panama
Acanthochitona andersoni, A.
balesae, A. rhodea, Acanthochites
rhodeus, Achanthochitona ferreirai:
6(1):79-114. Aequipectin circularis:
4(1):119. Calyptraea conica:
4(2):173-183. Cerithidea montagnei,
C. reevianum: 2:1-20. Crepidula
cerithicola, C. convexa, C. dilatata, C.
echinus, C. fecunda, C. incurva, C.
lessoni, C. plana, C. Striolata,
Crucibulum personatum, C.
scutellatum, C. spinosum, C. um-
brella: 4(2):173-183. Galeta Island:
6(1):79-114. Gatun Formation:
2:84-85; 4(1):1-12. Odostomia
(Chrysallida): 4(2):122. Ostrea ir-
ridescens: 4(1):119. Paleontology:
2:79, 84-85; 3(1):98. Pinctada
mazatlanica, Protothaca asperimma:
4(1):119. Turridae: 3(1):98. Turritella
abrupta: 4(1):1-12

Panamic Province
Paleontology: 2:84-85

Pascagoula River, MS
Corbicula fluminea: S2:7-39

Patuxent River, MD
Elliptio fisherianus, E. lanceolata, E.
productus: 3(1):94

Pauzar Rockshelter, KY
Abundance, Actinonaias ligamentina
carinata, Amblema plicata, Archae-
ology, Cyprogenia stegaria, Elimina
sp., Elliptio dilatata, Epioblasma
sampsoni, Fort Ancient People,
Fusconaia flava, F. maculata
maculata, Goniobasis sp., Human
Food, Kentucky, Kentucky River,
Lampsilis ventricosa, Lithasia
obovata, Magnonaias nervosa,
Obovaria retusa, O. subrotunda,
Physa sp., Pleurobema clava, P. cor-
datum, P plenum, P. rubrum, P sin-
toxia, Pleurocera canaliculatum,
Ptychobranchus fasciolaris,
Quadrula pustulosa, Q. quadrula:
1:31-34

Pea River, AL
Corbicula fluminea: S2:7-39

Peanut Island, FL
Acanthochitona andersoni, A.
balesae, A. roseojugum: 6(1):79-114

Pearl River, LA, MS
Corbicula fluminea: S2:7-39

Pearl River, PRC
Corbicula fluminalis: S2:113-124,
203-209. C. fluminea: S2:113-124

Peckerwood Creek, AL
Corbicula fluminea: S2:7-39

Pecos River, NM, TX
Corbicula fluminea: S2:7-39

276 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

Pee Dee River, SC
Corbicula fluminea: S2:7-39
Pennsylvania (PA)
Anodonta imbecilis: 4(2):231-232.
Corbicula fluminea: S2:7-39. Hender-
sonia occulta: 1:99. Leptoxis carinata:
4(1):119. Margaritifera margaritifera:
4(1):13-19. Pickering Creek:
4(2)231-232. Plagioporus hypentelli:
4(1):119. Susquehanna River: S2:7-39
Perdernales River, TX
Corbicula fluminea: S2:7-39
Persian Gulf
Cerithidea cingulata: 2:1-20
Peru
Acanthochitona rhodea: 6(1):79-114.
Chaco River: 3(1):96-97.
Eucrassatella gibbosa: 2:83.
Halodakra, H. (Halodakra) subtrigona:
3(1):103. Mollusca, unspecified,
Paleontology, Santa River: 3(1):96-97.
Turritella abrupta, Zoritos Formation:
4(1):1-12
Peruvian Province
Mollusca, unspecified: 3(1):96-97
Philippine Islands: 1:89
Cerithidea (Cerithdeopsilla), C.
cingulata: 2:1-20. Corbicula: S2:1-5.
C. fluminea, C. manilensis:
$2:113-124. Enigmonia aenigmatica,
Johohore Straits: 5(2):159-164.
Laguna de Bay, Luzon: S2:1-5.
Lepidozona (Lepidozona) luzonicus,
Luzon: 6(1):115-130. Masbate Island:
$1:23-34. Miocene, Negros Oriental:
2:1-20. Notoplax coarcata:
6(1):115-130. Perna viridis:
5(2):159-164. Pliocene: 2:1-20.
Quezon: 5(2):159-164. Solemya
(Acharax) bartschi, Tricas Island:
$1:23-34
Piankatank River, VA
Crassostrea virginica, Haplosporidium
nelsoni: S3:17-23
Pickering Creek, PA
Anodonta imbecilis: 4(2):231-232
Piney Creek, AL
Corbicula fluminea: S2:7-39
Piney Creek, TN
Corbicula fluminea: S2:7-39
Pinto Creek, TX
Corbicula: $2:125-132
Piscadera Baai, Curacao
Acanthochitona zebra: 6(1):79-114
Platte River, WI
Elliptio dilatatus delicatus:
5(2):165-171
Pocatalico River, WV
Corbicula fluminea: S2:7-39
Pokomoke Sound, VA
Crassostrea virginica, Haplosporidium
nelsoni: S3:17-23
Portugal
Corbicula fluminalis: S2:113-124

Potatoe Creek, GA
Corbicula fluminea: S2:7-39

Potatoe Slough, CA
Corbicula fluminea: S2:7-39

Potomac River, MD, VA, WV
Corbicula: S2:53-58. C. fluminea:
$2:7-39. Elliptio fisherianus, E.
lanceolata, E. productus: 3(1):94

Pound Creek, GA
Corbicula fluminea: S2:7-39

Powell River, TN
Actinonaias ligamentina, A. ligamen-
tina gibba, A. pectorosa, Alasmi-
donta marginata, Amblema plicata,
Cumberlandia monodonta,
Cyclonaias tuberculata tuberculata,
Cyprogenia stegaria, Dromus dromas
dromas, D. dromas caperatus,
Elliptio crassidens, E. dilatata,
Epioblasma brevidens, E. capsaefor-
mis, E. haysiana, E. lewisi, E.
torulosa gubernaculum, E. triquetra,
Fusconaia barnesiana, F. barnesiana
bigbyensis, F. cor. F. cor analoga, F.
cuneolus cuneolus, F. subrotunda, F.
subrotunda lesuerianus, Hemistena
lata, Lampsilis cardium, L. fasciola,
L. ovata, Lasmigona costata, L.
holstonia, Lemiox rimosa, Leptodea
fragilis, Lexingtonia dolabelloides,
Ligumia recta, L. recta latissima,
Medionidus conradicus, Plethobasus
cyphyus, Pleurobema oviforme, P.
oviforme argenteum, Potamilus
alatus, Ptychobranchus fasciolare, P.
subtentum, Quadrula cylindrica cylin-
drica, Q. cylindrica strigulata, Q. in-
termedia, Q. pustulosa, Q. sparsa,
Strophitus undulatus, Toxolasma
lividus lividus: 6(1):19-37. Unionids,
unspecified: 1:93-94. Villosa fabalis,
V. iris, V. vanuxemensis: 6(1):19-37

Poyang Lake, PRC
Corbicula largillierti: S2:113-124

Preston Rockshelter, WI
Amblema plicata, Anodonta grandis,
Anodontoides ferrussacianus, Elliptio
dilatata, E. dilatatus delicatus, Lamp-
silis radiata luteola, L. ventricosa,
Lasmigona complanata, Potamilus
alatus: 5(2):165-171

Prince Edward Island, Canada
Crassostrea virginica: S3:25-29

Puerto Rico (PR)
Acanthochiles (Notoplax) hemphilli,
Acanthochitona lineata, A. pygmaea,
A. zebra: 6(1):79-114. Biomphalaria
glabrata: 1:106, 1:107. Cayo Enrique,
Cryptoconchus floridanus, \sla Tur-
ramote, Magueyes Island, Media
Luna Reef: 6(1):79-114. Shistosoma
mansoni, S. mansoni Puerto Rican
PR-1, S. mansoni Puerto Rican
PR-2: 1:106. Solemya velum: S2:23-34

Punta Palmar, Yucatan, Mexico
Acanthochitona pygmaea: 6(1):79-114

Punta Rassa, FL
Acanthochitona pygmaea: 6(1):79-114

Qatar
Acanthochitona woodwardi, Chiton
peregrinus, C. (Rhyssoplax) affinis,
Ischnochiton winckworthi, |. yerbury,
Lepidozona luzonica, Notoplax
(Notoplax) arabica, Pinna muricata,
Tonicia (Lucilina) sueziensis:
6(1):115-130

Quaboag River, MA
Margaritifera margaritifera: 4(1):13-19

Queen River, RI
Margaritifera margaritifera: 4(1):13-19

Queensland, Australia
Berthella pellucida, Euselenops
luniceps, Moreton Bay, Pleuro-
branchus peronii: 5(2):197-214

Quintana Roo, Mexico
Acanthochitona andersoni, A.
Pygmaea: 6(1):79-114

Radley Pond, UK
Potamopyrgus jenkinsii: 5(1):73-84

Rappahannock River, VA
Crassostrea virginica: S3:17-23. Ellip-
tio fisherianus, E. lanceolata, E. pro-
ductus: 3(1):94. Haplosporidium
nelsoni: $3:17-23

Raritan River, NJ
Corbicula fluminea: 3(1):100-101;
$2:7-39

Red River, AR
Corbicula fluminea: S2:7-39

Red River, KY, TN
Actinonaias ligamentina gibba, A.
pectorosa, Alasmidonta marginata,
A. viridis, Amblema plicata perplicata:
6(1):19-37. Corbicula fluminea:
$2:7-39. Cyclonaias tuberculata,
Elliptio crassidens, E. dilatata,
Epioblasma florentina, E. florentina
walkeri, Lampsilis fasciola, L. ovata,
L. teres anodontoides, Lasmigona
complanata, L. costata, Megalonaias
nervosa, Obovaria retusa, Potamilus
alatus, Ptychobranchus fasciolare,
Strophitus undulatus, Tritogonia ver-
rucosa, Truncilla truncata, Villosa
vanuxemensis: 6(1):19-37

Red River, AR, LA, OK, TX
Corbicula fluminea: S2:7-39

Red Sea
Acanthopleura vaillantii: 6(1):115-130.
Anaspidea, Chicoreus virgineus:
4(1):109-110. Chiton huluensis, C.
(Rhyssoplax) affinis: 6(1):115-130.
Chromodoris africana: 5(2):243-258.
C. inornata, C. quadricolor Conidae,
Conus: 4(1):109-110. Cryptoplax
sykesi: 6(1):115-130. Elysia olivaceus,
Fasciolariidae: 4(1):109-110.
Gastropoda, unspecified: 4(1):103.

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 2h

Gymnodoris limaciformis:
4(1):109-110. Ischnochiton (Ischno-
chiton) yerburyi: 6(1):115-130. Murex
ramosus, Muricidae, Nerita forskali,
Neritidae: 4(1):109-110. Onithochiton
erythraeus: 6(1):115-130. Phyllida
varicosa, Phillodesmium xeniae,
Phyllobranchillus orientalis,
Pleuroploca trapezuim: 4(1):109-110.
Risbecia pulchella: 5(2):243-258.
Sacoglossa, Strombidae, Thaididae,
Thais savignyi: 4(1):109-110. Tonicia
(Lucilina) sueziensis: 6(1):115-130.
Trochidae, Trochus erythraeus, Tur-
binidae, Turbo radiatus: 4(1):109-110
Reelfoot Lake, TN
Amblema plicata, Anodonta grandis,
A. grandis corpulenta, A. imbecilis,
A. suborbiculata, Arcidens con-
fragosus, Lampsilis siliquoidea, Lep-
todea fragilis, Ligumia subrostrata,
Megalonaias nervosa, Plectomaris
dombeyanus, Quaarula pustulosa, Q.
quadrula, Toxolasma parva, T. tex-
asensis, Truncilla truncata: 6(1):19-37
Rhode Island (RI)
Amnicola limosa: 5(1):9-19. Arctica
islandica, Block Island: S3:51-57.
Campeloma decisum, Cipangopalu-
dina chinensis: 5(1):9-19. Crepidula
convexa, C. plana: 4(2):173-183. Fer-
rissia fragilis, F. parallela, Gyraulus
circumstriatus, G. deflectus, G. par-
vus, Helisoma anceps, H. cam-
panulatum, H. trivolvis, Laevapex
fuscus, Lyrogyrus granum, L.
pupoidea: 5(1):9-19. Margaritifera
margaritiferae: 4(1):13-19. Micromen-
tus dilatatus, Physa ancillaria, P
heterostropha, Planorbula armigera,
Promenetus exacuous, Pseudosuc-
cinea columella, Stagnicola elodes,
Valvata tricarinata, Viviparus georgi-
anus: 5(1):9-19
Rich Creek, TN
Corbicula fluminea: S2:7-39
Richardson Creek, NC
Corbicula fluminea: S2:7-39
Richland Creek, TN
Corbicula fluminea: S2:7-39
Ringtown Island, ME
Placopecten magellanicus: 6(1):1-8
Rio Grande, TX
Anodonta imbecilis henryiana, A.
grandis: 2:86. Corbicula fluminea:
2:86; S2:7-39. Cyrtonaias tampico-
ensis berlandieri, Disconaia
Salinasensis, Lampsilis teres,
Megalonaias gigantea, Popenaias
popei, Quadrula apiculata, Tox-
olasma parvus, Uniomerus
tetralasmus manubius: 2:86
Rio Grande do Sol, Brazil
Loligo sanpanulensis: 6(2):213-217

Riopel Pond, VA
Pisidium casertanum: 5(1):49-64
River Liffey, Republic of Ireland
Ancylus fluviatilis: 5(1):105-124
Roanoke River, NC
Anodonta implicata, Corbicula
fluminea, Elliptio complanata:
3(1):104-105
Roaring River, TN
Anodontoides ferussacianus, Lamp-
silis fasciola, Lasmigona costata,
Medionidus conradicus, Toxolasma
lividus glans, Villosa taeniata picta,
V. taeniata punctuata: 6(1):19-37
Roatan, Honduras
Acanthochiles (Notoplax) hemphilli,
Anthony Keys, Choneplax lata, Oak
Ridge: 6(1):79-114
Rockcastle River, TN
Corbicula fluminea: S2:7-39. Villosa
trabalis: 1:28
Rocky Creek, FL
Corbicula fluminea: S2:7-39
Rocky River, NC
Corbicula fluminea: S2:7-39
Roosevelt Lake, AZ
Corbicula fluminea, Ictiobus bubalus,
|. cyprinellus, |. niger: S2:7-39
Russian River, CA
Corbicula fluminea: S2:7-39
Rutherford Creek, TN
Corbicula fluminea: S2:7-39
Sabine River, LA, TX
Corbicula fluminea: S2:7-39
Sacramento River, CA
Corbicula: $2:125-132. C. fluminea:
4(1):81-89; S2:7-39, 133-142
St. Andrews Bay, FL
Acanthochitona pygmaea: 6(1):79-114
Croix River, MN, WI
Corbicula: S2:1-5. C. fluminea:
$2:7-39
Eustatius
Acanthochiton balesae, Tumble
Down Dick Bay: 6(1):79-114
St. Francis River, AR, MO
Corbicula fluminea: S2:7-39
Johns River, FL
Corbicula fluminea: S2:7-39. Elliptio
productus: 3(1):94
Joseph Bay, FL
Corbicula fluminea: S2:7-39.
Halodule wrightii, Laurencia poitei,
Marginella aureocincta, Rissoina
catesbyana, Thalassia testudinum:
4(2):185-199
Lucia
Acanthochitona andersoni:
6(1):79-114
Lucie Inlet, FL
Periploma margaritaceum: 2:35-40
Maarten
Acanthochitones spiculosus astriger:
6(1):79-114

St.

-

St.

-

St.

-

St.

-

St.

-

St.

-

St.

-

St. Marys Formation
Miliola marylandica, Teinostoma
nana: 4(1):39-42

St. Thomas, Virgin Islands
Acanthochitona pygmaea: 6(1):79-114

St. Vincent
Acanthochitona andersoni,
Choneplax lata: 6(1):79-114

Saipan
Cerithidea obtusa, Miocene: 2:1-20

Salinas River, CA
Corbicula fluminea: S2:7-39

Saline River, AR
Corbicula fluminea: S2:7-39

Saline River, IL
Corbicula fluminea: S2:7-39

Salkahatchie River, SC
Corbicula fluminea: S2:7-39

Salt Creek, IN
Corbicula fluminea: S2:7-39

Salt Pond, Bahama Islands
Acanthochitona zebra: 6(1):79-114

Salt River, AZ
Corbicula fluminea: S2:7-39

Salt River, KY
Corbicula fluminea: S2:7-39

Salton Sea, CA
Corbicula fluminea: S2:7-39

San Antonio, TX
Corbicula fluminea: S2:7-39

San Diego City Waterworks, CA
Corbicula fluminea: S2:7-39

San Francisco Bay, CA
Boccardia ligerica: S2:7-39.
Cerithidea californica: 2:1-20. Cor-
bicula fluminea, Corphium spinicoine,
C. stimpsoni, Macoma balthica:
S2:7-39

San Gabriel River, TX
Corbicula fluminea: S2:7-39

San Ignacio Formation, Baja California

Sur, Mexico
Amiantus sp., Calliostoma hannibali,
Chione (Chione) richthofeni, C.
(Chionopsis) sp., Choromytilus pallio-
punctatus, Crassilabrum wittichi,
Crassispira starri, Crepidula sp.,
Crucibulum inerme, C. scutellatum,
Cyclinellas, Cymia heimi, Cypraea
amandusi, Divalinga comis, Drillia
(Clathrodrillia) sp., Knefastia sp.,
Lucina (Lucinisca) sp., Macron hart-
mani, Mytilus canoasensis vidali,
Nassarius versicolor, Nerita
funiculata, Neverita (Glossaulax)
andersoni, Ostrea sp., Paleontology,
Sanguinolaria toulai, Solenosteira
sp., Strombina sp., Terebra burck-
hardti, Trachycardium sp., Turritella
bosei, T. costaricensis: 4(1):1-12

San Jacinto Reservoir, CA
Corbicula fluminea: S2:7-39

San Jacinto River, TX
Corbicula fluminea: S2:7-39

278 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

San Joaquin River, CA
Corbicula fluminea: S2:7-39, 133-142
San Juan Island, WA
Lepidochitona, L. dentiens: 4(2):243
San Luis Reservoir, CA
Corbicula fluminea: S2:7-39
San Nicolas Island, CA
Micrarionta opuntia: 3(1):98; 4(2):237.
M. sodalis: 3(1):98; 4(2):237
San-Men-Hsia Reservoir, PRC
Corbicula nitens: S2:113-124
Sand Key, FL
Acanthochiles (Notoplax) hemphilli:
6(1):79-114
Sandy Creek, MI
Corbicula fluminea: S2:7-39
Sangamon River, IL
Corbicula fluminea: S2:7-39
Sanibel Island, FL

Acanthochitona pygmaea: 6(1):79-114.

Anomia simplex, Argopecten gibbus,
Chione cancellata: 2:41-50
Santa Anna River, CA
Corbicula fluminea: S2:7-39
Santa Barbara Channel, CA
Cuthona albocrusta, Eubranchus,
Hermissenda crassicornis:
5(2):287-292
Santa Barbara Harbor, CA
Corbicula fluminea: S2:7-39
Santa Bouge Creek, AL
Corbicula fluminea: S2:7-39
Santa Catalina Island, CA
Opuntia littoralis, Orehelicidae,
Radiocentrum avalonense, Salvia
mellifera: 2:98
Santa Fe River, FL
Corbicula fluminea: S2:7-39
Santa River, Peru
Mollusca, unspecified: 3(1):96-97
Santee River, SC
Corbicula fluminea: S2:7-39
Sarasota Bay, FL
Acanthochitona pygmaea: 6(1):79-114
Saudi Arabi
Cerithidea cingulata: 2:1-20
Saugahatchee Creek, AL
Corbicula fluminea: S2:7-39
Savannah River, GA, SC
Corbicula: $2:1-5. C. fluminea:
$2:7-39. Crassostrea virginica:
$3:31-36
Scioto River, OH
Corbicula fluminea: S2:7-39
Scotland, UK
Acanthochitona crinita: 6(1):69-78.
Margaritifera margaritifera, Salmo
trutta: 5(1):125-128. Tonicella mar-
morea: 6(1):69-78
Sea of Galilee, Israel
Corbicula fluminalis: S2:113-124
Sebastian Inlet, FL
Ascobulla ulla, Elysia, E. ornata, Er-
colania funera, E. fuscovittata,

Lobiger souverbiei, Oxynoe antil-
larum, Placida: 5(2):259-280
Second Creek, AL
Corbicula fluminea: S2:7-39
Sepulga River, AL
Corbicula fluminea: S2:7-39
Sequatchie River, TN
Actinonaias pectorosa, Alasmidonta
viridus, Amblema plicata: 6(1):19-37.
Corbicula fluminea: S2:7-39.
Cumberlandia monodonta, Cyclon-
aias tuberculata, Elliptio crassidens,
E. dilatata, Epioblasma biemarginata,
Fusonaia barnesiana, Lampsilis
fasciola, Lasmigona costata, Lep-
todea fragilis, Obovaria subrotunda
lens, Pleurobema clava, Potamilus
alatus, Quadrula cylindrica, Tox-
olasma cylindrellus, Villosa iris, V.
vanuxemensis: 6(1):19-37
Seychelles
Baeolidida palythoae, Chromodoris,
C. africana, Haminoea natalensis,
Phyllidia, Pleurobranchus xhosa:
5(2):243-258. Tonicia (Lucilina) suez-
iensis: 6(1):115-130
Shanktown Creek, MS
Toxolasma texasensis, Unionmerus
tetralasmus: 4(1):21-23
Shasta Lake, CA
Corbicula fluminea: S2:7-39
Shoal Creek, TN
Corbicula fluminea: S2:7-39
Shubuta Creek, MS
Corbicula fluminea: S2:7-39
Sierra Nevada Mountains
Cooper, James Graham: 1:89
Siilaisenpuro River, Finland
Sphaerium corneum: 5(1):41-48
Silver Cove Canal, Bahama Islands
Acanthochitona lineata, A. worsfoldi,
A. zebra, Acanthochitones
spiculosus astriger: 6(1):79-114
Silver Creek, KY
Corbicula fluminea: S2:7-39
Singapore
Cerithidea cingulata: 2:1-20.
Lepidozona (Lepidozona) luzonicus:
6(1):115-130
Sinking Creek, TN
Corbicula fluminea: S2:7-39
Sister Creek, FL
Acanthochitona pygmaea: 6(1):79-114
Sky Lake, FL
Corbicula fluminea: S2:7-39
Slate Creek, KY
Corbicula fluminea: S2:7-39
Smith River, OR
Corbicula fluminea: S2:7-39
Snake River, ID, WA
Corbicula fluminea: S2:7-39
Solomon Islands
Maraunibina verrucosa, Paragantitus
ellynnae, Philinoglossa marcusi,

Pseudovermis mortoni, Pseudunela
cornuta: 5(2):281-286

Somalia
Chiton (Rhyssoplax) affinis,
Ischnochiton (Ischnochiton) yerburyi,
Tonicia (Lucilina) sueziensis:
6(1):115-130

Sonora, Mexico
Bulimulidae: 4(1):113-114. Hexaplex
erythrostomus, Muricanthus nigritus,
Octopus digueti: 6(1):45-48.
Orymaeus, Rhabdotus baileyi, R.
nigromontanus: 4(1):113-114

Souinlovey Creek, MS
Corbicula fluminea: S2:7-39

South Africa
Berthella plumula: 5(2):197-214

South America
Acanthochiles (Notoplax) hemphilli:
6(1):79-114. Angostura Formation:
4(1):1-12. Argentina: 2:21-34;
3(1):11-26; S2:1-5, 113-124. Biom-
phalaria glabrata, B. straminea, B.
tenagophila: 1:67-70. Brazil: 1:67-70,
92, 110; 2:21-34; 4(2):173-183, 233.
Buchanania onchidioides: 2:21-34.
Chaco River: 3(1):96-97. Chile:
2:21-34; 3(1):11-26. Colombia:
1:35-42; 4(1):1-12; 6(1):79-114. Cor-
bicula fluminea: S2:1-5, 113-124. C.
leana: S2:113-124. Crassatellinae:
2:83. Crassostrea rhizophorae:
1:35-42. Crepidula protea: 1:110;
4(2):173-183. Croton sp.-09: 1:67-70.
Ecuador: 2:84, 84; 3(1):98; 4(1):1-12;
$1:23-34. Esmereldas Formation:
2:84. Eucrassatella antillarum,
E. digueti, E. gibbosa: 2:83.
Fissurelidae annulus, Fissurella
patagonica, Fissurellidea megatrema,
F. patagonica: 2:21-34. Fusiturricula:
1:92. Halodakra, H. (Halodakra) sub-
trigona: 3(1):103. Littorina ziczac:
4(2):233. Mazatlania aciculata: 1:92.
Mollusca, unspecified: 2:84;
3(1):96-97. Neocorbicula: S2:113-124.
Paleontology: 3(1):96-97, 98;
4(1):1-12. Perna perna: 5(2):159-164.
Peru: 2:83; 3(1):96-97, 103. Pupillaea
annulus: 2:21-34. Santa River:
3(1):96-97. Siphonaria lessoni:
4(2):233. Solemya (Acharax)
johnsoni: S1:23-34. Trophon gever-
Sianus: 3(1):11-26. Turridae: 1:92;
3(1):98. Turritella abrupta, T. inezana,
T. ocoyana: 4(1):1-12. Uruguay:
2:21-34. Venezuela: 1:92; 2:83;
3(1):98

South Bay Aqueduct, CA
Corbicula fluminea: S2:7-39

South Bimini Island
Littorina mespillum, Puperita pupa,
Rissoella caribaea, Thalassia
testudinum: 4(2):185-199

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 21g

South Biscayne Bay, FL
Granulina ovuliformis, Halodule
wrightii, Laurencia poitei, Rissoinea
bryerea, Thalassia testudinum,
Tricolia affinis affinis: 4(2):185-199
South Carolina (SC)

Anadara brasiliana, A. broughtoni, A.

granosa, A. ovalis, A. transversa:
4(1):111. Clark Sound: 2:96-97;
4(2):149-155. Chione cancellata:
4(1):111. Congaree River: 1:95.
Cooper River, Corbicula fluminea:
S2:7-39. Busycon canaliculatum, B.
carica, B. contrarium, B. spiratum:
3(1):102. Edisto River: S2:7-39. Ellip-
tio angustata: 1:95. E. cistelliformis,
E. fisheriana, E. folliculata, E.
lanceolata, E. producta, E. ravenelli,
E. waccamawensis: 1:61-68. Hartwell
Reservoir, Intracoastal Waterway,
Lake Keowee: S2:7-39. Lampsilis
crocata, Leptodea ochracea: 1:61-68.
Little Pee Dee River: S2:7-39.
Mercenaria mercenaria: 2:96-97;
4(1):111; 4(2):149-155. Musculium par-
tumenium: S2:7-39. Noetia
ponderosa: 4(1):111. Octopus
vulgaris: 4(2):240. Pee Dee River:
$2:7-39. Polinices duplicatus:
4(1):111. Salkahatchie River, Santee
River, Savannah River: S2:7-39. Tox-
olasma pullus, Villosa ogeecheensis:
1:61-68. Waccamaw River: S2:7-39

South Chickamauga Creek, TN
Corbicula fluminea: S2:7-39

South Dakota (SD)
Anodonta grandis grandis, Lampsilis
teres teres, Lasmigona complanata,
Leptodea fragilis, L. leptodon,
Quadrula quadrula, Potamilus alatus,
P. ohiensis, Truncilla donaciformis, T.
truncata: 1:71-74

Spaanse Water, Curacao
Acanthochitona zebra: 6(1):79-114

Spring Creek, FL
Corbicula fluminea: S2:7-39

Spring Creek, TX
Corbicula fluminea: S2:7-39

Spring River, AR
Corbicula fluminea: S2:7-39

Sri Lanka (Ceylon)
Cancellaria lamellosa: 2:57-61.
Chiton huluensis, Dutch Bay:
6(1):115-130. Halgerda punctata:
§(2):243-258. Ischnochiton
(Ischnochiton) winckworthi, Notoplax
alisonae: 6(1):115-130. Trigonostoma
scalare: 2:57-61

Stanislaus River, CA
Corbicula fluminea: S2:7-39

Steel Bayou, MS
Corbicula fluminea: S2:7-39

Steinhatchie River, FL
Corbicula fluminea: S2:7-39

Stillwater River, OH
Corbicula fluminea: S2:7-39

Stones River, TN
Actinonaias ligamentina, A. pec-
torosa, Alasmidonta viridis, Amblema
plicata, Anodonta grandis, A. im-

becilis: 6(1):19-37. Corbicula fluminea:

§$2:7-39. Cumberlandia monodonta,
Cyclonaias tuberculata, Ellipsaria
lineolata, Elliptio dilatata, Epioblasma
arcaeformis, E. brevidens, E. floren-
tina, E. florentina walkeri, E. lenior,
Fusconaia flava, Lampsilis cardium,
L. fasciola, L. ovata, L. teres
anodontoides, Lasmigona com-
planata, L. costata, Leptodea fragilis,
Ligumia recta latissima, Medionidus
conradicus, Megalonaias nervosa,
Obliquaria reflexa, Obovaria sub-
rotunda, Pegias fabula, Pleurobema
coccineum, P. cordatum, P. oviforme,
P rubrum, Potamilus alatus, Ptycho-
branchus fasciolare, Quadrula cylin-
drica, Q. quadrula pustulosa, Q.
quaadrula, Simpsonaias ambigua,
Strophitus undulatus, Toxolasma
lividus, T. parva, Tritogonia verrucosa
Truncilla donaciformis, T. truncata:
6(1):19-37. Unionids, Unspecified:
1:93. Villosa iris, V. iris, V. lienosa, V.
trabalis: 6(1):19-37
Stoney Creek, IN
Corbicula fluminea: S2:7-39
Stow Lake, CA
Corbicula fluminea: S2:7-39
Strait of Juan de Fuca
Solemya agassizi: $1:23-34
Strait of Maccasar
Cancellaria lamellosa: 2:57-61
Strawberry River, AR
Corbicula fluminea: S2:7-39
Sucarnochee Creek, AL
Corbicula fluminea: S2:7-39
Suez Canal
Acanthopleura vaillanti, Chiton
huluensis: 6(1):115-130
Sugar Creek, TN
Corbicula fluminea: S2:7-39
Suislaw River, OR
Corbicula fluminea: S2:7-39
Sumatra, Indonesia
Cerithidea (Cerithdeopsilla): 2:1-20.
Corbicula gustaviana, C. moltkiana,
C. sumatrana, C. tobae, C. tumida:
$2:113-124. Java Sea, Lepidozona
(Lepidozona) luzonicus: 6(1):115-130.
Pliocene: 2:1-20
Sundu Sea
Moridilla brockii: 5(2):243-258
Sunflower River, MS
Corbicula fluminea: S2:7-39
Susquehanna River, MD, NY, PA
Corbicula fluminea: S2:7-39. Elliptio
productus: 3(1):94. Leptoxis carinata:
3(2):169-177

Suwanee River, FL
Campeloma geniculum: 3(1):99. Cor-
bicula fluminea: S2:7-39
Sweden
Embletonia pulchra: 5(2):303-306.
Sepietta oweniana: 2:90
Switzerland
Ancylus fluviatilis: 3(2):151-168
Tahiti
Durvilledoris leminiscata, Elysia
rufescens, Glossodoris atromar-
ginata, Gymnodoris ceylonica,
Hydatina amplustre, Pupa solidula:
5(2):243-258
Taiwan
Cerithidea (Cerithdeopsilla): 2:1-20.
Corbicula fluminea: S2:113-124.
Miocene: 2:1-20
Tallahalla Creek, MS
Corbicula fluminea: S2:7-39
Tallapoosa River, AL
Corbicula fluminea: S2:7-39
Tamarind Beach Reef, Bahamas
Acanthochitona andersoni, A.
pygmaea, A. zebra: 6(1):79-114
Tampa Bay, FL
Acanthochitona pygmaea:
6(1):79-114
Tangier Sound, VA
Crassostrea virginica, Haplosporid-
ium nelsoni: $3:17-23
Tanzania
Ceratophyllidia africana,
Chromodoris hamiltoni, C. vicina,
Cuthona kanga, Glossodoris, Jorun-
na zania, Sclerodoris coriacea:
5(2):243-258
Tar River, NC
Corbicula fluminea: 3(1):104-105.
Elliptio angustatus: 3(1):94. E. (Can-
thyria) steinstansana: 3(1):104-105.
E. emmonsi, E. fisherianus, E.
folliculatus, E. hazelhurstianus:
3(1):94. E. lanceolata: 1:94-95;
3(1):94. E. productus, E. shepar-
diana, E. subcylindraceus: 3(1):94
Tasman Sea
Chiton huluensis: 6(1):115-130
Tellico River, TN
Archaeology, Actinonaias ligamen-
tina, Anodonta grandis grandis, A.
imbecilis: 3(1):41-44. Corbicula
fluminea: 3(1):41-44; $2:7-39.
Dromus dromas, Elliptio crassidens,
E. dilatata, Epioblasma haysiana,
Fusconaia barnesiana, F. barne-
siana bigbyensis, F. subrotunda,
Lampsilis fasciola, L. ovata, Lex-
ingtonia dolabelloides, Medionidus
conradicus, Microyma nebulosa,
Pleurobema obliquum, P. oviforme,
P. oviforme argentium: 3(1):41-44.
Potamilus alatus: 3(1):41-44; 4(1):117.
Ptychobranchus subtentum,

280

Quaarula intermedia, Q. sparsa,
Strophitus undulatus, Toxolasma
lividus, Villosa iris, V. vanuxemensis:
3(1):41-44

Tennessee (TN)

Actinonaias carinata: 1:43-50;
6(1):19-37. A. carinata gibba:
6(1):19-37. A. ligamentina: 3(1):41-44;
4(1):25-37; 6(1):19-37; 6(2):165-178. A.
ligamentina gibba: 6(1):19-37. A. pec-
torosa: 1:43-50; 6(1):19-37.
Alasmidonta atropurpuata: 6(1):19-37.
A. calceolus: 1:43-50. A. marginata:
1:43-50; 6(1):19-37; 6(2):165-178. A.
minor: 1:43-50; 6(1):19-37. A.
raveneliana: 6(1):19-37. A. viridis:
6(1):19-37; 6(2):165-178. Amblema
costata: 1:43-50; 6(1):19-37. A.
costata perplicata, A. costata plicata:
6(1):19-37. A. peruviana: 6(1):19-37. A.
plicata: 1:43-50; 4(1):25-37, 117;
6(1):19-37; 6(2):165-178. A. plicata
perplicata, A. plicata plicata:
6(1):19-37. Anculosa praerosa:
1:43-50. Anodonta grandis: 1:43-50;
6(1):19-37; 6(2):165-178. A. grandis
corpulenta, A. grandis gigantea:
6(1):19-37. A. grandis grandis:
3(1):41-44; 6(1):19-37. A. imbecilis:
3(1):41-44; 6(1):19-37. A. suborbicu-
lata, Anodontoides ferussacianus, Ar-
cidens confragosus: 6(1):19-37. Bar-
ren Fork River, Big Bigby Creek, Big
Hickory Creek, Big Rock Creek:
$2:7-39. Big South Fork Cumberland
River: 6(1):19-37. Big Swann Creek:
$2:7-39. Buffalo River: 6(1):19-37;
$2:7-39. Busycon sp.: 4(1):25-37.
Campeloma sp.: 1:43-50; 4(1):25-37.
C. decisum: 6(2):165-178. Caney Fork
River: 4(1):117; 6(1):19-37. Carun-
culina glans: 6(1):19-37. C. lividus:
1:43-50. C. moesta, C. moesta cylin-
drella: 1:43-50; 6(1):19-37. C. parva,
C. texasensis: 6(1):19-37. Clinch
River: 4(1):25-37; 6(1):19-37; S2:7-39,
167-178. Collins River: S2:7-39. Con-
asauga River: 6(1):19-37. Conrdailla
caelata: 1:43-50; 4(1):25-37;
6(1):19-37. Corbicula fluminea:
3(1):41-44; 4(1):81-88; S2:7-39,
167-178. C. manilensis: 1:43-50.
Cumberland River: 4(1):81-88;
6(1):19-37; S2:7-39. Cumberlandia ir-
rorata: 4(1):25-37. C. mondonta:
4(1):13-19; 6(1):19-37. Cyclonaias
tuberculata: 1:43-50; 4(1):25-37;
6(1):19-37; 6(2):165-178. C. tuber-
culata granifera, C. tuberculata tuber-
culata, Cyprogenia irrorata:
6(1):19-37. C. stegaria: 4(1):25-37;
6(1):19-37; 6(2):165-178. Dromus
dromas: 1:43-50; 3(1):41-44;
4(1):25-37, 117; 6(1):19-37;

6(2):165-178. D. dromas caperatus, D.
dromas dromas: 6(1):19-37. Duck
River: 6(1):19-37; S2:7-39. Dysnomia
arcaeformis: 6(1):19-37. D. biemar-
ginata: 1:43-50. D. brevidens, D. cap-
saeformis: 1:43-50; 6(1):19-37. D. flex-
uosa: 6(1):19-37. D. florentina:
1:43-50; 6(1):19-37. D. florentina
walkeri: 6(1):19-37. D. haysiana:
1:43-50; 6(1):19-37. D. lenior, D.
lewisi, D. stewardsoni: 6(1):19-37. D.
torulosa: 1:43-50; 6(1):19-37. D.
torulosa gubernaculum, D. torulosa
propinqua: 6(1):19-37. D. triquetra:
1:43-50; 6(1):19-37. D. turgida:
6(1):19-37. East Rock River: S2:7-39.
Elk River: 1:43-50; 6(1):19-37;
$2:7-39. Elimia sp.: 4(1):25-37. Ellip-
saria lineolata: 6(1):19-37. Elliptio
crassidens: 1:43-50; 3(1):41-44;
4(1):25-37, 117; 6(1):19-37;
6(2):165-178. E. dilatata: 3(1):41-44;
4(1):25-37, 117; 6(1):19-37;
6(2):165-178. E. dilatatus: 1:43-50;
6(1):19-37. Emory River: 6(1):19-37;
$2:7-39. Epioblasma arcaeformis:
4(1):25-37; 6(1):19-37; 6(2):165-178. E.
biemarginata: 6(1):19-37. E.
brevidens, E. capsaeformis:
4(1):25-37; 6(1):19-37; 6(2):165-178. E.
flexuosa: 6(1):19-37. E. florentina:
4(1):117; 6(2):165-178. E. florentina
florentina, E. florentina walkeri:
6(1):19-37. E. haysiana: 3(1):41-44;
4(1):25-37; 6(1):19-37; 6(2):165-178. E.
lenior, E. lewisi: 6(1):19-37. E. obli-
quata: 4(1):25-37; 6(1):19-37. E. pro-
pinqua: 4(1):25-37; 6(1):19-37. E.
sampsoni: 1:27-30. E. stewardsoni:
4(1):25-37; 6(1):19-37; 6(2):165-178. E.
sulcata: 6(1):19-37. E. torulosa:
4(1):25-37; 6(1):19-37; 6(2):165-178. E.
torulosa cincinatiensis, E. torulosa
gubernaculum: 6(1):19-37. E. tri-
quetra: 4(1):25-37; 6(1):19-37. E.
turgidula: 6(1):19-37. Fall Creek, Flat
Creek, Fountain Creek: S2:7-39
French Broad River: 6(1):19-37.
Fusconaia barnesiana: 1:43-50;
3(1):41-44; 4(1):25-37; 6(1):19-37;
6(2):165-178. F barnesiana barne-
siana: 6(1):19-37. F. barnesiana big-
byensis: 1:43-50; 3(1):41-44;
6(1):19-37. F. barnesiana tumescens:
6(1):19-37. F cor analoga, F. cor cor:
6(1):19-37. F cuneolus: 4(1):43-50;
6(1):19-37. F. cuneolus appressa, F.
cuneolus cuneolus, F. ebena:
6(1):19-37. F. edgariana: 1:43-50;
6(1):19-37. F edgariana analoga, F.
flava, F. lateralis, F. polita, F. polita
lesueriana, F. polita pilaris, F. pusilla:
6(1):19-37. F subrotunda: 1:43-50;
3(1):41-44; 4(1):25-37, 117; 6(1):19-37;

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

6(2):165-178. F. subrotunda
leseuriana, F. subrotunda pilaris, F.
subrotunda subrotunda, F. undata:
6(1):19-37. Garrison River: S2:7-39.
Goniobasis laquetra: 1:43-50.
Greenlick Creek, Harpeth River,
Hatchie River: 6(1):19-37; S2:7-39.
Hemistena lata: 6(1):19-37;
6(2):165-178. Hendersonia occulta:
1:99. Hiwassee River: 6(1):19-37.
Holston River: 6(1):19-37; S2:7-39.
Horn Lake: 6(1):19-37. /o fluvialis:
4(1):25-37; 6(2):165-178. /. verrucosa
lima: 1:43-50. Lampsilis abrupta:
6(1):19-37. L. anodontoides: 1:43-50;
6(1):19-37. L. anodontoides fallaciosa,
L. cardium cardium, L. cardium
satura: 6(1):19-37. L. fasciola:
1:43-50; 3(1):41-44; 4(1):25-37;
6(1):19-37; 6(2):165-178. L. orbiculata:
4(1):25-37; 6(1):19-37. L. ovata:
1:43-50; 3(1):41-44; 4(1):25-37;
6(1):19-37; 6(2):165-178. L. ovata
satura: 6(1):19-37. L. ovata ven-
tricosa: 1:43-50; 6(1):19-37. L. sili-
quoida, L. teres, L. teres anodon-
toides: 6(1):19-37. L. teres teres:
4(1):117; 6(1):19-37. L. virescens:
6(1):19-37. Lasmigona badia:
6(1):19-37. L. complanata, L. costata:
1:43-50; 6(1):19-37; 6(2):165-178. L.
holstonia: 6(1):19-37; 6(2):165-178.
Lastena lata: 1:43-50; 6(1):19-37. Lep-
todea fragilis: 1:43-50; 6(1):19-37;
6(2):165-178. Lemiox rimosa:
4(1):25-37. L. rimosus: 6(1):19-37;
6(2):165-178. Leptodea leptodon:
6(1):19-37. Leptoxis (Athearnia)
crassa: 4(1):25-37. L. praerosa:
4(1):25-37; 6(2):165-178. Lexingtonia
dolabelloides: 1:43-50; 3(1):41-44;
4(1):25-37; 6(1):19-37; 6(2):165-178. L.
dolabelloides conradi: 1:43-50;
6(1):19-37. Lick River: S2:7-39.
Ligumia recta: 4(1):25-37, 117;
6(2):165-178. L. recta latissima, L.
subrostrata: 6(1):19-37. Lithasia
pinguis: 1:27-30. L. verrucosa:
4(1):25-37; 6(2):165-178. L. verrucosa
lima: 1:43-50. Little River: 6(1):19-37.
Little Duck River: S2:7-39. Little
Pigeon River, Little Pigeon River,
West Prong: 6(2):165-178. Little Ten-
nessee River: 6(1):19-37; S2:7-39.
Loosahgatchie River: 6(1):19-37.
McMahan Site: 6(2):165-178. Medi-
onidus conradicus: 1:43-50;
3(1):41-44; 6(1):19-37; 6(2):165-178.
Megalonaias gigantea: 1:43-50;
6(1):19-37. M. nervosa: 4(1):117;
6(1):19-37. Microyma nebulosa:
3(1):41-44. Mississippi River:
6(1):19-37; S2:7-39. Nine Mile Creek:
$2:7-39. Nolichucky River: 6(1):19-37;

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 281

S2:7-39. North Fork Creek: S2:7-39.
North Fork Obion River: 6(1):19-37.
Notchy Creek: S2:7-39. Obey River:

6(1):19-37; S2:7-39. Obliquaria reflexa:

1:43-50; 6(1):19-37. Obovaria olivaria,
O. retusa: 6(1):19-37. O. subrotunda:
1:43-50; 6(1):19-37; 6(2):165-178. O.
subrotunda lens: 1:43-50; 4(1):25-37;
6(1):19-37. O. subrotunda levigata:
6(1):19-37. Paint Rock River: S2:7-39.
Pegias fabula: 1:43-50; 6(1):19-37.
Piney River: S2:7-39. Plagiola
lineolata: 1:43-50; 6(1):19-37. Pletho-
basus cicatricosus: 4(1):25-37;
6(1):19-37. P cooperianus, P
cyphyus: 4(1):25-37; 6(1):19-37;
6(2):165-178. P. cyphyus compterus,
P. pachosteus, P. striatus: 6(1):19-37.
Pleurobema aldrichianum: 6(1):19-37.
P. clava: 4(1):25-37; 6(1):19-37. P
clava catillus, P coccineum:
6(1):19-37. P cordatum: 1:43-50;
4(1):25-37; 6(1):19-37; 6(2):165-178. P
gibberum, P. obliquata: 6(1):19-37. P
obliquum: 3(1):41-44; 6(1):19-37. P
oviforme: 1:43-50; 3(1):41-44;
6(1):19-37; 6(2):165-178. P. oviforme
argentium: 1:43-50; 3(1):41-44;
6(1):19-37. PR. oviforme holstonense, P.
permorsa: 6(1):19-37. P. plenum:
1:27-30; 4(1):117; 6(1):19-37;
6(2):165-178. P. rubrum: 6(1):19-37;
6(2):165-178. Pleurocera
canaliculatum: 1:43-50; 4(1):25-37;
6(2):165-178. P. canaliculatum un-
dulatum: 4(1):25-37. P_ parvum:
6(2):165-178. Potamilus alatus:
3(1):41-44; 4(1):117; 6(1):19-37;
6(2):165-178. P. ohioensis: 6(1):19-37.
Powell River: 6(1):19-37. Proptera
alata: 1:43-50; 6(1):19-37. P
laevissima: 6(1):19-37. Ptycho-
branchus fasciolare: 6(1):19-37. P
fasciolaris: 1:43-50; 4(1):25-37;
6(2):165-178. P. subtentum: 1:43-50;
3(1):41-44; 4(1):25-37; 6(1):19-37;
6(2):165-178. Quadrula bullata:
6(1):19-37. Q. cylindrica: 1:43-50;
4(1):25-37; 6(1):19-37; 6(2):165-178. Q.
cylindrica strigillata: 6(1):19-37. Q.
fragosa: 6(1):19-37. Q. intermedia:
1:43-50; 3(1):41-44; 4(1):25-37;
6(1):19-37. Q. metanevra: 1:43-50;
4(1):25-37; 6(1):19-37. Q. nodulata:
6(1):19-37. Q. pustulosa: 1:43-50;
4(1):25-37; 6(1):19-37; 6(2):165-178. Q.
quaorula: 1:43-50; 6(1):19-37. Q.
sparsa: 3(1):41-44; 4(1):19-37;
6(2):165-178. Red River: 6(1):19-37;
$2:7-39. Reelfoot Lake: 6(1):19-37.
Rich Creek, Richland Creek:
$2:7-39. Roaring River: 6(1):19-37.
Rutherford Creek: S2:7-39.

Sequatchie River: 6(1):19-37; S2:7-39.

Shoal Creek: S2:7-39. Simpsoni-
concha ambigua, Simpsonaias am-
bigua: 6(1):19-37. Sinking Creek,
South Chickamauga Creek: S2:7-39.
Stones River: 1:93; 6(1):19-37;
S2:7-39. Strophitus rugosus: 1:43-50;
6(1):19-37. Strophitus undulatus
1:43-50; 3(1):41-44; 6(1):19-37. Sugar
Creek: S2:7-39. Tellico River:
3(1):41-44; S2:7-39. Tennessee River:
4(1):25-37; 6(1):19-37; S2:7-39. Tox-
olasma cylindrella, T. livida:
6(1):19-37. T. lividus: 3(1):41-44;
6(2):165-178. T. lividus glans, T.
lividus lividus, T. parva, T. texasensis:
6(1):19-37. Tritogonia verrucosa, Trun-
cilla donaciformis, T. truncata:
1:43-50; 6(1):19-37. T. vermiculata,
Uniomerus tetralasmus: 6(1):19-37.
Unionids, Unspecified: 1:93. Villosa
fabalis: 1:43-50; 6(1):19-37. V. iris:
1:43-50; 3(1):41-44; 6(1):19-37;
6(2):165-178. V. lienosa: 6(1):19-37. V.
nebulosa: 1:43-50; 6(1):19-37. V. per-
purpurea: 6(1):19-37. V. picta:
6(1):19-37. V. taeniata: 1:43-50;
4(1):25-37; 6(1):19-37. V. taeniata pic-
ta, V. taeniata punctata, V. taeniata
taeniata, V. teneltus: 6(1):19-37. V.
trabalis: 1:27-30; 4(1):25-37;
6(1):19-37; 6(2):165-178. V. trabalis
perpurpurea: 6(1):19-37. V. vanuxemi:
1:43-50, 4(1):25-37. V. vanuxemensis:
3(1):41-44; 6(1):19-37; 6(2):165-178.
Watauga River: 6(1):19-37. Watts Bar
Reservoir: S2:167-178. Weekly Creek:
$2:7-39. Wolf River: 6(1):19-37

Tennessee Reef, FL

Acanthochitona andersoni, A. zebra:
6(1):79-114

Tennessee River, AL, KY, TN

Actinonaias ligamentina, A. ligamen-
tina gibba, A. pectorosa, Alasmidon-
ta marginata, A. viridis, Amblema
plicata, A. plicata plicata, Anodonta
grandis, A. grandis corpulenta, A.
imbecilis, A. suborbiculata, Acidens
confragosus: 6(1):19-37. Corbicula
fluminea: S2:1-5, 7-39. Cumberlandia
monodonta, Cyclonaias tuberculata
tuberculata, C. tuberculata granifera,
Cyprogenia stegaria, Dromus dromas
dromas, D. dromas caperatus, Ellip-
saria lineolata, Elliptio crassidens, E.
dilatata, E. dilatata subgibbosus,
Epioblasma arcaeformis, E.
biemarginata, E. brevidens, E. cap-
saeformis, E. flexuosa, E. florentina,
E. florentina walkeri, E. haysiana, E.
lenior, E. lewisi, E. obliquata, E. pro-
pinqua, E. stewartsoni, E. torulosa
torulosa, E. torulosa gubernaculum,
E. triquetra, E. turgidula, Fusconaia
barnesiana barnesiana, F. barnesiana

bigbyensis, F. barnesiana tumescens,
F. cor, F. cor analoga, F. cor cor, F.
cuneolus appressa, F. cuneolus
cuneolus, F. ebena, E. flava, F. flava
trigona, F. subrotunda, F. subrotunda
lesuerianus, F. subrotunda pilaris,
Hemistena lata, Lampsilis abrupta, L.
cardium, L. fasciola, L. ovata, L.
teres anodontoides, L. teres teres, L.
virescens, Lasmigona complanata, L.
costata, L. holstonia, Lemiox
rimosus, Leptodea fragilis, L. lep-
todon, Lexingtonia dolabelloides,

L. dolabelloides conradi, Ligumia
recta, L. recta latissima, Medi-
onidus conradicus, Megalonaias
nervosa, Obliquaria reflexa, Obovaria
olivaria, O. retusa, O. subrotunda, O.
subrotunda levigata, O. subrotunda
lens, Pegias fabula, Plethobasus
cicatricosus, P cooperianus, P
cyphyus, P cyphyus compterus,
Pleurobema catillus, P clava, P. coc-
cineum, P. cordatum, P. oviforme, P
oviforme argenteum, P. oviforme
holstonse: 6(1):19-37. P. plenum:
1:27-30; 6(1):19-37. P rubrum,
Potamilus alatus, P ohioensis,
Ptychobranchus fasciolare, P subten-
tum, Quadrula cylindrica cylindrica,
Q. cylindrica strigulata, Q. fragosa,
Q. intermedia, Q. metanevra, Q.
nodulata, Q. pustulosa, Q. quadrula,
Q. sparsa, Strophitus undulatus, Tox-
olasma cylindrellus, T. lividus glans,
T. lividus lividus, T. parva, Tritogonia
verrucosa, Truncilia donaciformis, T.
truncata, Uniomerus tetralasmus,
Villosa fabalis, V. iris, V. taeniata
picta, V. taeniata taeniata, V. trabalis,
V. trabalis perpurpurea, V. vanux-
emensis: 6(1):19-37

Tensas River, LA

Corbicula fluminea: S2:7-39

Terrapin Creek, AL

Corbicula fluminea: S2:7-39

Texas (TX)

Angelina River: S2:7-39. Anodonta
imbecilis henryiana: 2:86. A. grandis:
2:86; S2:179-184. Aplocinotus grun-
niens.: S2:7-39. Benbrook Lake:
S$2:179-184. Big Cypress River,
Blanco River: S2:7-39. Bradley
Creek, Bradley Reservoir:
$2:179-184. Brazos River: S2:7-39,
179-184. Calliostoma roseolum, C.
velieli: 2:84. Cedar Creek Reservoir:
$2:179-184. Clear Fork, Trinity River:
$2:151-166. Colorado River: S2:7-39,
125-132. Compano Bay: 1:89.
Concho River: S2:7-39, 179-184. Cor-
bicula: S2:125-132. C. fluminea: 2:86;
$2:7-39, 99-111, 151-166, 179-184,
193-201, 231-239. Crassostrea

282 AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988

virginica: S3:25-29. Cyrtonaias
tampicoensis berlandieri, Disconaia
salinasensis: 2:86. Elm Fork, Trinity
River: S2:179-184. Falcon Reservoir:
2:86. Gastropoda, Freshwater,
Unspecified, Gastropoda, Terrestrial,
Unspecified: 1:99. Guadalupe River:
S2:7-39, 179-184. Johnson Creek:
S2:7-39. Laguna Madre: 1:89. Lake
Arlington: 3(2):267-268; S2:99-111,
231-239. Lake Fairfield: S2:125-132.
Lake Long: S2:179-184. Lake of the
Pines: S2:125-132. Lake Raven:
$2:179-184. Lake Theo: 1:99;
$2:179-184. Lampsilis teres: 2:86.
Lepomis microphus: S2:7-39.
Lewisville Lake: S2:179-184. Little
Brazos River: S2:7-39. Llano Grande
Lake: S2:179-184. Llano River:
$2:7-39, 179-184, 193-201. Lysinoe:
3(1):102-103. Matagorda Bay:
2:35-40. Megalonaias gigantea: 2:86.
Melampus bidentatus: 4(1):121-122;
4(2):236. Minytrema melanops,

Nueces River: S2:7-39. Onion Creek:

$2:179-184. Paleontology:
3(1):102-103. Pecos River, Perder-
nales River: S2:7-39. Periploma
margaritaceum, P. orbiculare:
2:35-40. Physella virgata:
3(2):243-265. Pinto Creek:
$2:125-132. Popenaias popei,
Quaarula apiculata: 2:86. Q.
quaarula: S2:99-111 (passim). Rana
catesbeiana: S2:179-184. Red River:
$2:7-39. Rio Grande: 2:86; S2:7-39.
Sabine River, San Antonio River,
San Gabriel River, San Jacinto
River, Spring Creek: S2:7-39. Tox-
olasma parvus: 2:86. Trinity River:
$2:7-39; 151-166, 179-184. Twin
Buttes Reservoir: S2:179-184.
Uniomerus tetralasmus manubius:
2:86. White River: S2:7-39

Thailand
Cerithidea cingulata, C. obtusa, C.
quadrata: 1:20. Corbicula arata, C.
baudoni, C. blandiana, C. bocourti,
C. erosa, C. fluminea, C. guber-
natoria, C. gustaviana, C. heardi, C.
iravadica, C. javanica, C. lamarck-
iana, C. larnaudieri, C. leviuscula, C.
ligidiana, C. lydigiana, C. messageri,
C. moreletiana, C. noetlingi, C. oc-
cidentiformis, C. petiti, C. pingensis,
C. pisidiformis, C. regia, C. siamen-
sis, C. solidula, C. tenuis, C.
virescens, C. vokesi: S2:113-124.
Perna viridis: 5(2):159-164

Thomas Hill Reservoir, MO
Corbicula fluminea: S2:7-39

Tibbee Creek, MS
Corbicula fluminea: S2:7-39

Timor, Indonesia
Corbicula australis: S2:113-124

Timor Sea
Chiton huluensis: 6(1):115-130
Tioughnioga River, NY
Leptoxis carinata: 3(2):169-177
Tobago
Choneplax lata: 6(1):79-114
Tolumne River, CA
Corbicula fluminea: S2:7-39
Tombigbee River, AL, MS
Corbicula fluminea: S2:7-39
Torres Straits
Chiton huluensis: 6(1):115-130
Tortuga Island, Venezuela
Acanthochitona andersoni, A.
balesae: 6(1):79-114
Towaliga River, GA
Corbicula fluminea: S2:7-39
Town Creek, AL
Corbicula fluminea: S2:7-39
Tradewater River, KY
Corbicula fluminea: S2:7-39
Tred Avon River, MD
Crassostrea virginica: S3:25-29
Trinidad
Acanthochiton balesae, Avalon Bay:
6(1):79-114. Paleontology, Turridae:
3(1):98
Trinity River, TX
Clear Fork: S2:151-166. Corbicula
fluminea: S2:7-39, 151-166
Trout Lake, WI
Amnicola limosa, Campeloma
decisa, Ferrissia, Haemopsis gran-
dis, Lepomis gibbosus, L.
microlophus, Leucochloridismorpha
constantine, Lymnaea elodes, L.

emarginata, L. stagnalis, Umbra limi:

5(1):73-84
Tuamoto Archipelago
Pleurehdera haraldi: 5(2):197-214
Tubbs Creek, AL
Corbicula fluminea: S2:7-39
Tumble Down Dick Bay, St. Eustatius
Acanthochiton balesae: 6(1):79-114
Tung-ting Lake, PRC
Corbicula largillierti, C. nitens:
$2:113-124
Tully Lake, NY
Viviparous georgianus: 3(2):268
Tunisia
Bulinus truncatus, Hydrobia aponen-
sis, Melanoides tuberculata,
Melanopsis, Mercuria confusa, M.
punica: 5(1):85-90
Turks and Caicos Islands
Acanthochiles (Notoplax) hemphilli,
Acanthochitona pygmaea, Crypto-
conchus floridanus: 6(1):79-114
Twelve Pole Creek, WV
Corbicula fluminea: S2:7-39
Twin Buttes Reservoir, TX
Corbicula fluminea: S2:179-184
Tygarts Creek, KY
Corbicula fluminea: S2:7-39

Uchee Creek, AL
Corbicula fluminea: S2:7-39

Uhwarrie River, NC
Corbicula fluminea: S2:7-39

Umpqua River, OR
Corbicula fluminea: S2:7-39

Unadilla River, NY
Leptoxis carinata: 3(2):169-177

Union of Soviet Socialist Republics

(USSR)

Ancylus fluviatilis, Caspian Sea:
3(2):151-168. Corbicula cor, C.
ferganensis, C. fluminalis, C. fluminea,
C. japonica, C. purpurea, C. tibeten-
sis: $2:113-124

United Arab Emirates
Acanthopleura vaillantii, Chiton
peregrinus, Ischnochiton winckworthi,
Lepidozona luzonica: 6(1):115-130

United Kingdom
Ancylus fluviatilis: 3(2):151-168. Arch-
idoris pseudoargus: 4(1):103-104.
Biomphalaria glabrata, Bulinus jous-
seaumei: 5(1):65-72. Embletonia
pulchra: 5(2):303-306. Eukiefferiella
sp.: 3(2):151-168. Margaritifera
margaritifera: 5(1):125-128. Mytilus
edulis, M. galloprovincialis: 1:108.
Northern Ireland: 5(2):303-306. Physa
fontinalis, Planorbis planorbis, P
vortex: 5(1):65-72. Potamopyrgus
jenkinsii, Radley Pond: 5(1):73-84.
Salmo trutta, Scotland: 5(1):125-128.
Theba pisana: 1:104

Uruguay
Fissurellidea megatrema: 2:21-35

Utla Island, Bahama Islands
Acanthochitona roseojugum:
6(1):79-114

Vaca Key, FL
Acanthochitona balesae, A.
pygmaea, Cryptoconchus floridanus:
6(1):79-114

Venezuela
Acanthochitona andersoni, A.
balesae, A. rhodea, A. venezuelana:
6(1):79-114. Eucrassatella antillarum:
2:83. Isla be Margarita: 6(1):79-114.
Mazatlania aciculata: 1:92. Paleon-
tology, Turridae: 3(1):98. Tortuga
Island: 6(1):79-114

Verde River, AZ
Corbicula fluminea: S2:7-39

Virgin Islands
Acanthochitona lineata, A. pygmaea,
Choneplax lata, St. Thomas:
6(1):79-114

Virginia (VA)
Acteocina canaliculata, Acteon
wetherilli: 4(1):39-42. Actinonaias
pectorosa, Alasmidonta marginata,
A. minor: 3(1):104. A. viridis,
Ambloplites rupestris, Anadonta
anatina: 5(1):1-7. Appomattox River: |

AMER. MALAC. BULL. GEOGRAPHIC INDEX: 1983 - 1988 283

$2:7-39. Balanus concavus, B.
finchii, B. proteus: 4(1):39-42. Big
Moccasin Creek: 5(1):1-7. Bivalvia,
unspecified: 4(2):231. Campostoma
anomalum: 5(1):1-7. Chesapeake
Bay: 2:79; S3:17-23. Chickahominy
River: S2:7-39. Clinch River:
4(2):231; S2:7-39. Columbellidae:
3(1):96. Concavus finchii, Conus
marylandicus: 4(1):39-42. Corbicula:
$2:1-5, 53-58. C. fluminea: 4(1):116;
5(1):1-7; S2:7-39, 69-81. Cottus
carolinae: 5(1):1-7. Coyner Springs:
3(1):99-100. Crassostrea virginica:
$3:17-23, 31-36. Crepidula costata:
4(1):39-42. Dugesia tigrina: S2:7-39.
Elizabeth River: S3:31-36. Elliptio
fisherianus, E. lanceolata, E. produc-
tus: 3(1):94. Etheostoma flabellare, E.
rufilineatum: 5(1):1-7. Farriers Pond:
5(1):49-64. Fusconaia barnesiana:
3(1):104; 5(1):1-7. F edgariana:
3(1):104. Fusinus pumilus: 4(1):39-42.
Goniobasis proxima: 3(1):99-100.
Great Wicomico River, Haplosporidium
nelsoni: S3:17-23. Hendersonia
occulta: 1:99. Holston River, North
Fork: 3(1):104. James River: 3(1):94;
$2:7-39; S3:17-23, 31-36. Juliamitrella:
3(1):96. Lampsilis fasciola: 3(1):104;
6(1):1-7. L. ovata, Lasmigona costata:
3(1):104. L. subviridis: 6(2):179-188.
Lexingtonia dolabelloides: 3(1):104.
Mactra clathrodon, M. modicella, M.
subcuneata: 4(1):39-42. Medionidus
conradicus: 3(1):104; 5(1):1-7;
6(2):179-188. Micropterus dolomieui:
5(1):1-7. Milford Haven: S3:17-23.
Miliola marylandica, Mitrella com-
munis: 4(1):39-42. Mobjack Bay:
$3:17-23. Mulinia lateralis: 4(1):39-42.
New River: 4(1):116; S2:1-5, 7-39,
69-81. Nocomis micropogon, Notrop-
sis coccogenis, N. galacturus:
5(1):1-7. Odostomia (Chesapeakella):
3(1):96. Oenopota pumilus:
4(1):39-42. Paleontology: 2:79;
3(1):96; 4(1):39-42. Pelecypoda: 2:79.
Piankatank River: S3:17-23. Pisidium
casertanum: 5(1):1-7, 49-64. P com-
pressum: 5(1):1-7. Pleurobema
oviforme: 3(1):104; 5(1):1-7;
6(2):179-188. Pocomoke Sound:
$3:17-23. Potomac River: 3(1):94;
$2:7-39, 53-58. Ptychobranchus
fasciolaris, P subtentum: 3(1):104.
Pyramidellidae: 3(1):96. Quin-
queloculina semiluna: 4(1):39-42.
Rappahannock River: 3(1):94;
$3:17-23. Riopel Pond: 5(1):49-64.
Rotella nana: 4(1):39-42. Sphaerium
striatinum: 5(1):1-7. Spisula confraga,
S. modicella: 4(1):39-42. Tangier
Sound: $3:17-23. Teinostoma nana:

4(1):39-42. Toxolasma lividus:
3(1):104. Utriculastra: 4(1):39-42.
Villosa nebulosa: 3(1):104; 5(1):1-7. V.
vanuxemi: 3(1):104; 5(1):1-7;
6(2):179-188. York River: S3:17-23

Virginia Key, FL
Alvania auberiana, Caecum nitidum,
Halodule wrightii, Laurencia poitei,
Rissoina bryerea, Smaragdia viridis
viridemaris, Thalassia testudinum:
4(2):185-199

Virginian Province
Extinction, Faunal Replacement,
Paleontology: 2:79

Viscaino Peninsula, Mexico
Paleontology: 2:84-85

Wabash River, IL, IN
Corbicula fluminea: S2:7-39.
Epioblasma sampsoni, Quadrula
cylindrica, Strophitus undulatus: 1:28

Waccamaw River, NC, SC
Corbicula fluminea: S2:7-39

Waccassa River, FL
Corbicula fluminea: S2:7-39

Water Island
Acanthochitona lineata: 6(1):79-114

Watts Bar Reservoir, TN
Corbicula fluminea: S2:167-178

Washington (WA)
Acochlidacea: 2:95. Archidoris
montereyensis: 4(2):205-216.
Chehalis River, Columbia River:
$2:7-39. Cooper, James Graham:
1:89. Corbicula fluminea: S2:7-39.
Lepidochitona, L. dentiens: 4(2):243.
Nucella lamellosa: 3(1):11-26.
Pseudovermis: 2:95. Snake River:
$2:7-39

Watauga River, TN
Actinonaias pectorosa, Amblema
marginata, Elliptio dilatata, Fusconaia
barnesiana bigbyensis, F. subrotun-
da, F. subrotunda lesuerianus, Lamp-
silis fasciola, L. ovata, Lasmigona
costata, Leptodea fragilis, Medi-
onidus conradicus, Pleurobema
oviforme argenteum, Strophitus un-
dulatus, Villosa iris, V. vanuxemensis:
6(1):19-37

Wateree River, NC
Corbicula: $2:125-132

Weekly Creek, TN
Corbicula fluminea: S2:7-39

Wekiva River, FL
Corbicula: S2:1-5. C. fluminea:
$2:7-39

West Africa
Pluerobranchus tarda: 5(2):243-258

West Drain, NM
Corbicula fluminea: S2:7-39

West Fork River, WV
Corbicula fluminea: S2:7-39

West Hawksbill Creek, Grand

Bahama Island

Acanthochitona pygmaea: 6(1):79-114
West Indies
Acanthochitones spiculosus:
6(1):79-114. Voluta cancellaria:
2:57-61
West Summerland Key, FL
Corbicula fluminea: S2:7-39. Epio-
blasma sampsoni: 1:28
White River, TX
Corbicula fluminea: S2:7-39
Whitewater River, MO
Corbicula fluminea: S2:7-39
Wicomico River, MD
Corbicula fluminea: S2:7-39
Willamette River, OR
Corbicula fluminea: S2:7-39
Wisconsin (WI)
Actinonaias ligamentina carinata:
1:51-60; 5(2):165-171. Alasmidonta
marginata: 1:51-60; 5(2):165-171. A.
viridis, Amblema plicata:
5(2):165-171. A. plicata plicata:
1:51-60. Amnicola limosa: 5(1):73-84.
Anodonta grandis: 5(2):165-171. A.
grandis corpulenta, A. grandis
grandis, A. imbecilis, A. suborbicu-
lata: 1:51-60. Anodontoides ferrussa-
cianus: 5(2):165-171. Arcidens con-
fragosus: 1:51-60; 5(2):165-171.
Brogley Rockshelter: 5(2):165-171.
Campeloma decisa: 5(1):73-84. Cor-
bicula fluminea: S2:7-39. Cyclonaias
tuberculata, Ellipsaria lineolata:
1:51-60. Elliptio crassidens
crassidens: 1:51-60; 5(2):165-171. E.
dilatata: 1:51-60; 5(2):165-171.
E. dilatatus delicatus: 5(2):165-171.
Ferrissia: 5(1):73-84. Fusconaia
ebena: 1:51-60; 5(2):165-171. F. flava:
1:51-60; 5(2):165-171. Grant River:
5(2):165-171. Haemopsis grandis:
5(1):73-84. Hendersonia occulta:
1:51-60. Lampsilis higginsi: 1:51-60;
4(2):230. L. radiata luteola: 1:51-60;
5(2):165-171. L. teres anodontoides:
1:51-60; 5(2):165-171. L. teres teres:
1:51-60; 5(2):165-171. L. ventricosa:
1:51-60; 5(2):165-171. Lasmigona
complanata: 1:51-60; 5(2):165-171. L.
compressa: 5(2):165-171. L. costata:
1:51-60; 5(2):165-171. Lepomis gib-
bosus, L. microlophus: 5(1):73-84.
Leptodea fragilis: 1:51-60.
Leucochloridismorpha constantine:
5(1):73-84. Ligumia recta: 1:51-60;
5(2):165-171. Little Grant River:
5(2):165-171. Lymnaea elodes, L.
emarginata, L. stagnalis: 5(1):73-84.
Magnonaias nervosa: 1:51-60;
5(2):165-171. Millville Site:
5(2):165-171. Mississippi River:
4(2):230; 5(2):165-171.. Obovaria
Olivaria: 1:51-60. Platte River:
5(2):165-171. Plethobasus cyphyus,

284 AMER. MALAC. BULL. GEOGRAPHIC INDEX:

Pleurobema rubrum, P. sintoxia:
1:51-60. Potamilus alatus: 1:51-60;
5(2):165-171. P. ohiensis: 1:51-60.
Preston Rockshelter: 5(2):165-171.
Quaarula metanerva, Q. nodulata:
1:51-60. Q. pustulosa: 5(2):165-171.

Q. pustulosa pustulosa, Q. quadrula:

1:51-60. Strophitus undulatus un-
dulatus: 1:51-60; 5(2):165-171. Tox-

olasma parvus, Tritogonia verrucosa:

1:51-60. Trout Lake: 5(1):73-84. Trun-
cilla donaciformis, T. truncata:
1:51-60. St. Croix River: S2:7-39.
Strophitus undulatus undulatus:
5(2):165-171. Umbra limi: 5(1):73-84.
Venustaconcha ellipsiformis ellipsi-
formis, Villosa iris iris, Wisconsin
River: 5(2):165-171

Wisconsin River, WI
Arcidens confragosus, Fusconaia
ebena, F. flava, Lampsilis teres
anodontoides: 5(2):165-171

Withlacoochee River, FL, GA
Corbicula fluminea: S2:7-39

Wolf River, TN
Anodonta suborbiculata, Lampsilis
teres teres, Leptodea fragilis,

Potamilus purpurata, Quadrula

pustulosa mortoni, Tritogonia ver-

rucosa, Truncilla: 6(1):19-37
Woodford River, Republic of Ireland

Ancylus fluviatilis: 5(1):105-124
Woods Hole, MA

Chaetopleura apiculata: 6(1):69-78.

Crepidula convexa, C. fornicata, C.

plana, Limulus polyphemus, Littorina

littorea, Lunatia heros: 3(1):33-40
Woodward Creek, MS

Corbicula fluminea: S2:7-39
Yalobusha River, MS

Corbicula fluminea: S2:7-39
Yangtze River, PRC

Corbicula largillierti: S2:113-124
Yazoo River, MS

Corbicula fluminea: S2:7-39
Yellow River, FL

Corbicula fluminea: S2:7-39
Yellow River, PRC

Corbicula nitens: S2:113-124
Yemen

Acanthopleura vaillantii: 6(1):115-130
Yockanookany River, MS

Corbicula fluminea: S2:7-39

1983 - 1988

Yucatan Peninsula
Acanthochitona pygmaea: 6(1):79-114.
Crassostrea rhizophorae, C. virginica:
1:108. Punta Palmar: 6(1):79-114
York River, VA
Crassostrea virginica, Haplosporidium
nelsoni: $3:17-23
Yorktown Formation
Conus marylandicus, Crepidula
costata, Miliola marylandica,
Oenopota pumilus, Spisula confraga,
Teinostoma nana: 4(1):39-42
Yucun Balam, Mexico
Acanthochitona pgymaea: 6(1):79-114
Yugoslavia
Lepidopleurus cajetanus: 6(1):153-159
Zanzibar
Chiton (Chiton) fosteri, Ischnochiton
(Ischnochiton) yerburyi: 6(1):115-130
Zimbabwe
Biomphalaria glabrata: 1:106-107. B.
pfeifferi: 5(1):85-90. B. straminea,
Bulinus natalensis, B. tropicus, B.
truncata, Mazoe Dam: 1:106-107
Zorritos Formation, Peru
Paleontology: 4(1):1-12

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 285

Acetate Peel

Lasmigona subviridis, Medionidus
conradicus, Pleurobema oviforme,
Villosa vanuxemi: 6(2):179-188

Acid Rain

Amnicola limosa, Anodonta grandis
corpulenta, Campeloma decisum,
Cincinnatia concinnatiensis, Corbicula
fluminea, Elliptio complanata,
Gyraulus parvus, Helisoma anceps,
Lampsilis radiata, Musculium securis,
Physella gyrina, Pisidium spp., P
variable, Sphaerium spp., Valvata
tricarinata: 5(1):31-39.

Adaptation

Aciculidae: 3(2):223-231. Acochlidi-
acea: 5(2):281-286. Adalaria proxima:
4(1):103-104. Ampullariidae:
3(2):223-231. Aplacophora:
5(2):281-286. Archidoris pseudoargus:
4(1):103-104. Assimineidae, Bithynii-
dae, Buccinum undatum:
3(2):223-231. Cadlina laevis:
4(1):103-104. Caecum: 5(2):281-286.
Cerithiidae: 3(2):223-231. Corbicula
fluminea: S2:223-229. Cyclophori-
dae, Deroceras reticulatum:
3(2):223-231. Eupera cubensis:
$2:223-229. Gastrohedyle:
5(2):281-286. Haliotis corrugata, H.
rufescens: 3(2):223-231. Hedylopsis:
5(2):281-286. Helicinidae:
3(2):223-231. Helisoma trivolvis:
3(2):243-265. Helix pomatia, Hydrobi-
idae, Hydrocenidae: 3(2):223-231.
Jorunna tormentosa: 4(1):103-104.
Limax pseudoflavus, Littorina irrorata:
3(2):223-231. Lymnaea (Stagnicola)
elodes: 3(2):143-150. L. stagnalis:
3(2):223-231. Maraunibina verrucosa:
5(2):281-286. Marisa cornuarietis:
3(2):223-231. Meiomenia,
Meiopriapulus fijiensis: 5(2):281-286.
Melaniidae, Melanoposidae, Meso-
gastropoda: 3(2):223-231. Musculium
spp.: S2:223-229. Neomeniomorpha:
5(2):281-286. Neopisidium:
$2:223-229. Nerita fulgurans,
Neritacea, Neritidae, Neritina
latissima: 3(2):223-231. Nudi-
branchia: 5(2):281-286. Onchidoris
muricata: 4(1):103-104. Opistho-
branchia, Paraganitus ellynnae:
5(2):281-286. Patella vulgata:
3(2):223-231. Philinoglossa, P._ mar-
cusi: 5(2):281-286. Pisidium spp.:
$2:223-229. Pleuroceridae, Pomacea
lineata, Potamopyrgus jenkinsii:
3(2):223-231. Pseudovermis spp.,
Pseudunela, P cornuta: 5(2):281-286.

SUBJECT INDEX

Rissoacea, Rissoidae: 3(2):223-231.
Sphaerium spp.: S2:223-229. Strom-
bus gigas: Syrnolopsidae, Thiaridae:
3(2):223-231. Tritonia hombergi:
4(1):103-104. Valvatacea, Valvatidae,
Viviparacea, Viviparidae, Viviparus
spp.: 3(2):223-231

Adaptation, Shape

Lasaeidae: 1:90. Leptonacea:
1:90-91

Adductor Muscle

Lasmigona costata: 2:82

Aerial Exposure

Crepidula convexa spp.: 3(1):33-40.
Polymesoda caroliniana: 6(2):199-206

Aesthetes

Age

Lepidopleurus cjetanus: 6(1):153-159

Biomphalaria glabrata: 1:106. Cor-
bicula fluminea: S2:151-166. Illex il-
lecebrosus: 4(2):240-241. Lasmigona
subviridis: 6(2):179-188. Leptoxis
carinata: 3(2):169-177. Medionidus
conradicus, Pleurobema oviforme:
6(2):179-188. Spirodon carinata:
3(2):169-177. Villosa vanuxemi:
6(2):179-188

Agglutin, Human Anti-A

Pulmonata: 1:97-98

Aging Techniques

Fusconaia barnesiana, Pleurobema
oviforme: 3(1):106

Alimentary System

Cerithidea scalariformis: 2:1-20

Allometry

Cistopus indicus: 6(2):207-211. Ellip-

tio icterina: 1:95. Transennella tantilla:

2:94. Hapalochlaena maculosa,
Octopus spp., Pteroctopus tetra-
cirrhus, Robsonella mfontanianus,
Scaeurgus patagiatus, S. unicirrhus:
6(2):207-211. Villosa villosa: 1:95

Allozymes

Acahtina fulica: 6(1):16. Adalaria pro-
xima: 6(1):7. Amblemini: 1:109-110.
Anguspira alternata: 6(1):16. Arion
spp.: 1:110; 6(1):16. Austrocohlea
constricta, Bathybembix bairdi:
6(1):17. Bradybaena similaris: 6(1):16.
Biomphalaria spp., Campeloma
geniculum, C. parthenum: 6(1):17.
Cepaea spp., Cerion bendalli, C. in-
canum: 6(1):16. Cerithium spp.:
6(1):17. Corbicula: 1:96; S2:125-132.
Crassostrea spp.: 1:108, 109.
Crepidula spp.: 1:110; 6(1)17.
Deroceras spp.: 1:110; 6(1):16. Elliptio
spp., Elliptoideus, Fusconaia:
1:109-110. Goniobasis spp.: 6(1):17.
G. proxima: 1:105. Helisoma trivolvsis,

Helix aspersa, H. pomatia: 6(1):16.
Lampsilis: 1:109-110. Liguus spp.:
5(2):153-157. Limax spp.: 6(1):16. Lit-
torina arcana, L. rudis, Lymnaea
elodes, Melanoides tuberculata:
6(1):17. Mercenaria mercenaria: 1:107.
Mesodon zaletus: 2:97-98. Millax
budapestensis, M. gagates, M.
sowerbyi: 6(1):16. Nassarius obsoleta:
6(1):17. Nymphophilus minckleyi:
6(1):16. Onchidoris muricata: 6(1):17.
Otala lactea, Oxychillas cellarius,
Partula spp.: 6(1):16. Physa hetero-
stropha: 6(1):17. Pisidium casertanum:
5(1):49-64. Potamopyrgus jenkinsi:
6(1):17. Quadrula, Quincuncina:
1:109-110. Rumina decollata, Sphinc-
terochila spp.: 6(1):16. Thais
haemastoma, T. lamellosa: 6(1):17.
Theba pisana: 6(1):16. Triodopsis:
2:97-98. T. albolabris: 6(1):16.
Viviparous contectoides: 6(1):17.
Xerocrassa saetzeni: 6(1):16

Anatomy

Acanthochiton fascicularis:
6(1):141-151. Alaba, Alba goniochila:
4(2):235. Aplacophora: 6(1):57-68.
Ashmunella chiricahuna: 2:98.
Bulinus tropicus: 1:96. Calyptraeidae,
Calyptraea conica, C. mamillaris, C.
novazelandiae: 4(2):173-183.
Cerithiidae: 4(2):235. Chaetopleura
lurida, C. peruviana: 6(1):141-151.
Chiton olivaceus: 6(1):131-139,
141-151. Corbicula fluminea: 1:13-20;
3(1):101; S2:113-124, 223-229. C.
spp.: S2:113-124. Crepidula spp.,
Crucibulum spp.: 4(2):173-183. Diala
goniochila, Diastomidae: 4(2):235.
Elliptio angustata, E. lanceolata:
1:95. Eudoxochiton nobilis:
6(1):141-151. Eupleura caudata
etterea: 2:63-73. Fissurellidea spp.:
2:21-34. Helminthoglyptidae: 2:98.
Hipponix grayanus: 4(2):173-183.
Ischnochiton herdmani, Katharina
tunicata, Lepidochitona cinerea, L.
dentiens, Lepidozona retiporosus,
Lepidopleurus cajetanus: 6(1):141-151.
Litiopa, Litiopidae: 4(2):235.
Megatebennus spp.: 2:21-34.
Mesodon elevatus: 2:98. Mesodon
zaletus: 1:98. Monadenia fidelis:
2:98. Mopalia spp.: 6(1):141-151. Of
fadesma angasi: 2:35-40. Orthalicus
spp.: 2:98. Perna viridis:
5(2):159-164. Placiphorella velata:
6(1):141-151. Planaxidae: 4(2):235.
Planaxis: 2:1-20. Plaxiphora obtecta:
6(1):141-151. Pleioptygma,

P. helenae: 3(1):97-98. Polygyridae:
2:98. Polyplacophora: 6(1):57-68.
Pupillaea spp.: 2:21-34. Sonorella
virilis: 2:98. Thais haemastoma
canaliculata: 2:63-73. Thracia
pubescens: 2:35-40. Tonicella in-
signis: 6(1):141-151. Triodopsis spp.:
1:98. Urosalpinx cinerea, U. cinerea
follyensis: 2:63-73

Anatomy, Comparative

Acado: 5(2):215-241. Acanthopleura
granulata: $1:1-22. Aciculidae:
3(2):223-231. Aclididae, Aclis,
Acochlidiacea, Acteocina sp.,
Acteocinidae, Acteon: $1:1-22.
Adalaria spp.: 2:95. Aeolidacea:
5(2):215-241. Aglaja, Aglajidae,
Akera, Akeridae, Allogastropda,
Amaea, Amphibola, Amphibolidae:
$1:1-22. Ampullariidae: 3(2):223-231.
Anaspidea, Angutispira: S1:1-22.
Anidolyta, A. spongotheras:
5(2):215-241. Anodonta spp.:
4(1):13-19. Anthobranchia:
5(2):215-241. Aplysia sp., Aplysiidae,
Aplysiomorpha, Architectonicacea:
$1:1-22. Arminacea : 5(2):215-241.
Ascoglossa: $1:1-22. Assimineidae:
3(2):223-231. Atyidae: $1:1-22.
Austrophon: 3(1):11-26. Basomma-
tophora: $1:1-22. Bathyberthella
spp.: 5(2):215-241. Batillaria spp.,
Batillariinae: 2:1-20. Berthelinia:
$1:1-22. Berthella spp.: 5(2):215-241;
$1:1-22. Berthellina citrina, B. engeli,

Berthellinae, Birthellini, Berthellinops:

5(2):215-241. Bithyniidae:
3(2):223-231. Bittum: 2:1-20.
Blauneria: $1:1-22. Boonea: $1:1-22.
Boreotrophon spp.: 3(1):11-26. Buc-
cinacea: 3(1):11-26. Buccinum un-
datum: 3(2):223-231. Buchanania
onchidioides: 2:21-34. Bulla: S1:1-22.
B. membranacea, B. plumula:
5(2):215-241. Bullidae, Bullina,
Bullomorpha, Calyptraeidae:
$1:1-22. Campanile, Campanilidae,
Carychium, Cephalaspidea: $1:1-22.
Cerithiacea, Cerithidea spp.,
Cerithideopsis: 2:1-20. Cerithiidae:
2:1-20; 3(2):223-231. Cerithiopsacea,
Cerithiopsidae: S1:1-22. Cerithium
spp.: 2:1-20. Chelidonura: S1:1-22.
Chicoreus palmarosae: 3(1):11-26.
Chilina, Chilinidae: $1:1-22. Clado-
branchia, Cleanthus: 5(2):215-241.

Couthouyella: $1:1-22. Cumberlandia,

C. monodonta: 4(1):13-19.
Cyanogaster: 5(2):215-241. Cyclo-
phoridae: 3(2):223-231; S1:1-22.
Cyclostremella, Cyclostremellidae,
Cylinchna, Cylindrobulla, Cylin-
drobullidae, Cymbulia, Cymbuliidae,

Cylindrobulla: S1:1-22. Dendronota-
cea: 5(2):215-241. Deroceras
reticulatum: 3(2):223-231. Detracia,
Diaphana, Diaphanidae,
Diaphorodoris: 2:95. Docoglossa:
$1:1-22. Doridacea: 5(2)215-241.
Ebala, Ellobiidae, Ellobium, Elysia,
Elysiidae, Entomotaeniata,
Epitoniacea, Epitoniidae, Epitonium,
Eulimacea, Eulimidae: $1:1-22.
Eupera cubensis: S2:223-229.
Euselenops, E. luniceps:
5(2):215-241. Euthyneura, Fargoa
bartschi: S1:1-22. Gastroplax:
5(2):215-241. Gegania: $1:1-22.
Gigantonotum: 5(2):215-241. Gleba:
$1:1-22. Gourmya gourmyi: 2:1-20.
Gymnosomata: S1:1-22. Gym-
notoplax, G. americanus:
5(2):215-241. Haliotis corrugata, H.
rufescens: 3(2):223-231. Haminoea,
Hedylopsidae, Hedylopsis,
Heliaucus, H. cylindricus, H. per-
reieri: S1:1-22. Helicinidae, Helix
pomatia: 3(2):223-231. Hetero-
branchia, Heterogastropoda,

Heteroglossa, Hydatina, Hydatinidae:

$1:1-22. Hydrobiidae, Hydrocenidae:
3(2):223-231. Janthina sp., J. exigua,
J. janthina, Janthinidae: S$1:1-22.
Joannisia: 5(2):215-241. Juliidae:
$1:1-22. Koonsia: 5(2):215-241.
Lamellidens, Lampsilis, L. radiata:
4(1):13-19. Latia, Latiidae,
Leucophytia, Limacinidae, Limapon-
tia, Limapontiidae: $1:1-22. Limax
pseudoflavus: 3(2):223-231. Lirularia,
L. lirulata: 4(1):109. Littorina: $1:1-22.
L. irrorata: 3(2)223-231. Lymacina:
$1:1-22. Lymnaea stagnalis:
3(2):223-231. Macfarlandaea:
5(2):215-241. Magilidae: 3(1):11-26.
Margaritifera margaritifera, M. mar-
rianae, Margaritiferidae: 4(1):13-19.
Marinula: $1:1-22. Marisa cor-
nuarietis: 3(2):223-231. Mathilda,
Mathildidae, Maxacteon, Melam-
pidae, Melampus: S$1:1-22.
Melaniidae, Melanoposidae:
3(2):223-231. Melanopsis: 2:1-20.
Mesogastropoda: 3(2):223-231;
$1:1-22. Micromelo: $1:1-22.
Modulus: 2:1-20. Murex acantho-
stephes, Muriciacea: 3(1):11-26.
Muricidae: 3(1):11-26; S1:1-22.
Muricopsinae: 3(1):11-26. Musculium
spp.: S2:223-229. Myxa: $1:1-22.
Neda: 5(2):215-241. Neogastropoda:
$1:1-22. Neopisidium: S2:223-229.
Neotrigonia sp.: 4(1):13-19. Nerita
fulgurans, Neritacea, Neritidae,
Neritina latissima: 3(2):223-231.
Notaspidea: 5(2):215-241; $1:1-22.

286 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Nucella lamellosa: 3(1):11-26. Nudi-
branchia: S1:1-22. Oceanebridae,
Odontocymbiolinae: 3(1):11-26.
Odostomia, Omalogyra: $1:1-22. Om-
brella: 5(2):215-241. Onchidella,
Onchidiidae, Onchidium: $1:1-22.
Onchidoris spp.: 2:95. Operculatum:
5(2):215-241. Opisthobranchia:
$1:1-22. Oscaniopsis, Oscaniella,
Oscanius: 5(2):215-241. Otina,
Otinidae, Ovatella, Oxynidae,
Oxynoe: $1:1-22. Parmophorus,
Patella perversa, P umbraculum:
5(2):215-241. P vulgata: 3(2):223-231.
Paziella, P pazi: 3(1):11-26. Percale,
Peraclidae, Phanerophthalmus,
Philine, Philinidae, Philinoglossa,
Philinoglossidae, Philippa: S1:1-22.
Pisidium spp.: S2:223-229. Planorbi-
dae: S1:1-22. Pleurehdera, P. haraldi,
Pleurobranchacea, Pleurobranchaea,
P maculata, P meckelii, Pleuro-
branchaeidae, Pleurobranchella, P
alba, P. nicobarica: 5(2):215-241.
Pleurobranchidae: 5(2):215-241;
$1:1-22. Pleurobranchidium, Pleuro-
branchillus, Pleurobranchinae,
Pleurobranchoides gilchristi:
5(2):215-241. Pleurobranchomorpha:
$1:1-22. Pleurobranchus spp.:
5(2):215-241; S1:1-22. Pleuroceridae:
3(2):223-231. Poirieri: 3(1):11-26.
Pomacea lineata: 3(2):223-231.
Potamides spp., Potamididae,
Potamidinae: 2:1-20. Potamopyrgus
Jenkinsii: 3(2):223-231. Prosobranchia,
Pseudomalaxis, Pseudoskenella,
Ptenoglossa, Pulmonata, Pupa, Pur-
pura patula, Pyramidella crenulata,
Pyramidellacea, Pyramidellidae:
$1:1-22. Pyrazus, P ebininus: 2:1-20.
Pythia: $1:1-22. Rachiglossa:
3(1):11-26. Radix, Retusidae,
Retussa: $1:1-22. Rhinoclava (Pro-
clava) kochii: 2:1-20. Ringicula,
Ringiculidae: S1:1-22. Rissoacea:
3(2):223-231. Rissoella, Rissoellidae:
$1:1-22. Rissoidae: 3(2):223-231.
Roxania: $1:1-22. Roya, R.
spongotheras: 5(2):215-241. Saco-
glossa, Salinator Sayella,
Scaphander, Scaphandridae:
$1:1-22. Siphonaria: 5(2):215-241;
$1:1-22. Siphonariidae, Smarag-
dinella: $1:1-22. Sphaerium spp.:
$2:223-229. Spiricella: 5(2):215-241.
Stiligar, Stiligeridae: S1:1-22. Strom-
bus gigas: 3(2):223-231. Susania:
5(2):215-241. Syrnolopsidae:
3(2):223-231. Systellommatophor:
$1:1-22. Telescopium, Terebralia, T.
palustris: 2:1-20. Thais haemastoma,
T. lapillus: 2:63-73. Thecosomata:

AMER.

$1:1-22. Thiaridae: 3(2):223-231.
Toledonia, Triopohridae, Trochidae:
$1:1-22. Trophon spp., Trophoninae:
3(1):11-26. Turbonilla, T. vineae, Tur-
ritellidae: S1:1-22. Tylodina spp.,
Tylodinella, T. trinchesii, Tylodindae:

5(2):215-241. Tympanotonus fascatus:

2:1-20. Umbonium: 4(1):109. Um-
braculacea: 5(2):215-241. Umbraculi-
dae, Umbraculum: 5(2):215-241;
$1:1-22. U. umbraculum, Umbrella:
5(2):215-241. Valvata: $1:1-22.
Valvatacea, Valvatidae: 3(2):223-231;
$1:1-22. Velesunio: 4(1):13-19. Veroni-
cellidae: S1:1-22. Viviparacea,
Viviparidae, Viviparus spp.:

3(2):223-231. Volvatella, Volvatellidae:

$1:1-22. Williamia: 5(2):215-241;
$1:1-22. Zaccatrophon: 3(1):11-26.
Zemelanopsis: 2:1-20. Zidoninae:
3(1):111-26

Anatomy, Demibranch
Elliptio lanceolata: 1:94-95

Anatomy, Reproductive
Ashmunella chiricahuna, Mesodon
zaletus, Stenotrema fraternum,
Triodopsis albolabris: 1:98

Anoxia
Pisidium amnicum, P. personatum,
Sphaerium corneum, S. transversum
(passim): 5(1):41-48

Aposematic Coloration
Opisthobranchia: 5(2):185-196,
243-258, 287-292

Aquaculture
Aequipectin circularis: 4(1):119. Cor-
bicula fluminea: S2:211-218.
Mercenaria mercenaria: 4(2):149-155.
Ostrea irridescens: 4(1):119. Perna
viridis: 5(2):159-164 (passim). Pinc-
tade mazatlanica, Protothaca
asperimma: 4(1):119.

Aquarium Display
Loligo opalescens, Nautilus pom-
pilius, Octopus dolfleini, O.
rubescens, Sepia officinalis: 4(2):241

Aragonite
Byssus: 2:41-50

Archaeology
Actinonaias ligamentina: 3(1):41-45;
4(1):25-37; 6(2):165-178. A. ligamen-
tina carinata: 1:31-34; 5(2):165-171.
Alasmidonta marginata, A. viridis:
5(2):165-171; 6(2):165-178. Amblema
plicata: 1:31-34; 4(1):25-37;
5(2):165-171; 6(2):165-178. Anodonta
grandis: 5(1):91-99; 6(2):165-178. A.
grandis corpulenta, Anodontoides

ferussacianus, Arcidens confragosus:

5(2):165-171. Busycon sp.,
Campeloma sp.: 4(1):25-37. C.
decisum: 6(2):165-178. Conradiilla
caelata, Cumberlandia monodonta:

4(1):25-37. Cyclonaias tuberculata:
4(1):25-37; 6(2):165-178. Cyprogenia
irrorata: 4(1):25-37. C. stegaria:
1:31-34; 4(1):25-37; 6(2):165-178.
Dallas Component, McMahon Site,
TN: 6(2):165-178. Dromus dromas:
3(1):41-45; 4(1):25-37; 6(2):165-178.
Elimia sp.: 4(1):25-37. Elimina sp.:
1:31-34. Elliptio crassidens:
4(1):25-37; 6(2):165-178. E.
crassidens crassidens: 5(2):165-171.
E. dilatata: 1:31-34; 4(1):25-37;
5(2):165-171; 6(2):165-178. E. dilatatus
delicatus: 5(2):165-171. Epioblasma
spp.: 1:31-34; 3(1):41-45; 4(1):25-37;
6(2):165-178. Fort Ancient People:
1:31-34. Fusconaia barnesiana:
4(1):25-37; 6(2):165-178. F ebena:
5(2):165-171. F. flava: 1:31-34;
5(2):165-171. F maculata maculata:
1:31-34. F subrotunda: 3(1):41-45;
4(1):25-37. Goniobasis sp.: 1:31-34.
Hemistena lata: 6(2):165-178. lo
fluvialis: 4(1):25-37; 6(2):165-178.
Lampsilis fasciola: 4(1):25-37;
6(2):165-178. L. orbiculata: 4(1):25-37.
L. ovata: 4(1):25-37; 6(2):165-178. L.
spp.: 5(2):165-171. L. ventricosas:
1:31-34; 5(2):165-171. Lasmigona
complanata, L. compressa:
5(2):165-171. L. costata: 4(1):25-37;
5(2):165-171; 6(2):165-178. L.
holstonia: 6(2):165-178. Lemiox
rimosa: 4(1):25-37. L. rimosus:
6(2):165-178. Leptodea fragilis:
4(1):25-37; 5(2):165-171. Leptoxis
(Athearnia) crassa: 4(1):25-37. L.
praerosa: 4(1):25-37; 6(2):165-178.
Lexingtonia dolabelloides: 3(1):41-45;
4(1):25-37; 6(2):165-178. Ligumia rec-
ta: 4(1):25-37; 5(2):165-171;
6(2):165-178. Lithasia (Angitrema)
verrucosa: 6(2):165-178. L. geniculata
salebrosa: 4(1):25-37. L. obovata:
1:31-34. L. verrucosa: 4(1):25-37.
Magnonaias nervosa: 1:31-34. Medi-
onidus conradicus: 3(1):41-45;
6(2):165-178. M. nervosa:
5(2):165-171. Obovaria retusa:
1:31-34; 4(1):25-37. O. subrotunda:
1:31-34; 6(2):165-178. O. subrotunda
lens: 4(1):25-37. Pauzar Rockshelter,
KY, Physa sp.: 1:31-34. Plethobasus
cicatricosus: 4(1):25-37. P
cooperianus: 4(1):25-37; 6(2):165-178.
P. cyphyus: 4(1):25-37; 5(2):165-171;
6(2):165-178. Pleurobema clava:
1:31-34; 4(1):25-37. P cordatum:
1:31-34; 4(1):25-37; 6(2):165-178. P
obliqum: 3(1):41-44. P. oviforme:
3(1):41-44; 6(2):165-178. P plenum, P
rubrum: 1:31-34; 6(2):165-178. P. sin-
toxia: 1:31-34. Pleurocera

MALAC. BULL. SUBJECT INDEX: 1983 - 1988 287

canaliculatum: 1:31-34; 4(1):25-37;
6(2):165-178. P. canaliculatum un-
dulatum: 4(1):25-37. P parvum:
6(2):165-178. Potamilus alatus:
5(2):165-171; 6(2):165-178. Ptycho-
branchus fasciolaris: 1:31-34;
4(1):25-37; 6(2):165-178. P. subtentum:
3(1):41-45; 4(1):25-37; 6(2):165-178.
Quadrula cylindrica: 4(1):25-37;
6(2):165-178. Q. intermedia: 3(1):41-45;
4(1):25-37. Q. metanerva: 4(1):25-37;
5(2):165-171. Q. pustulosa: 1:31-34;
4(1):25-37; 5(2):165-171; 6(2):165-178. Q.
quaadrula: 1:31-34; 5(2):165-171. Q. spar-
Sa. 3(1):41-45; 6(2):165-178. Strophitus
undulatus undulatus: 5(2):165-171. Tox-
olasma lividus: 6(2):165-178. Tritogonia
verrucosa, Venustaconcha ellipsiformis
ellipsiformis: 5(2):165-171. Villosa spp.,
3(1):41-45; 4(1):25-37; 5(2):165-171;
6(2):165-178
Arm Suckers
Cephalopoda: 6(2):207-211
Attachment
Anomia simplex, Chlamys islandica:
$1:35-50. Crassostrea virginica:
$3:41-49. Mytilus edulis, Ostrea
edulis, Patella vulgata, Pecten max-
imus, Unela nahantensis: $1:35-50
Batesian Mimicry
Opisthobranchia: 5(2):185-196, 287-292
Behavior
Abra alba: 5(1):21-30. Achatina fulica:
2:98-99. Acmaeia scabra: S1:35-50.
Aegires sublaevis, Aeolidia papillosa,
Aeolidiella glauca, A. sanguinea,
Aeolidiopsis, Aldisa, A. banyulensis:
5(2):185-196. Anatina papyratia: 2:35-40.
Anisodoris: 5(2):185-196. Aplysia spp.:
2:78; 5(2):185-196. Archidoris spp.:
5(2):185-196. Astarte castanea:
5(1):21-30 (passim). Ataagena:
5(2):185-196. Brechites penis: 5(1):21-30
(passim). Bursatella: 5(2):185-196.
Calocochlea, C. caillaudi: 3(1):98-99.
Cardiomya planetica: 1:13 (passim).
Catriona gymnota: 5(2):185-196. Chiton
olivaceus: 6(1):131-139. Chlamys oper-
cularis: 1:13 (passim). Cochlodesma
praetenue: 2:35-40; S1:35-50.
Cochlostyla (Hypselostyla) carinata, C.
(Orthostylus) pithogaster, C.
pithogaster: 3(1):98-99. Collembola:
5(2):185-196. Collisella scabra: S1:35-50.
Corbicula: 5(1):21-30 (passim);
$2:41-45. C. fluminea: 1:13-20;
4(1):61-79, 81-88; S1:35-50, 187-191,
193-201. Corbiculacea: 5(1):21-30
(passim). Coryphella: 5(2):185-196.
Crassostrea virginica: 4(1):101; S3:41-49.
Crepidula spp.: 3(1):33-40. Cuthona
spp., Dendrodoris, Discodoris, Dondice
paguerensis, Dolabrifera, Doridella

288 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

obscura, Doridella steinbbergae,
Doridomorpha gardineri, Doriopsilla,
D. pharpa, Doris, Elysia arena,
Eubranchus exiguus, Facelina cor-
onata, Favorinus branchialis:
5(2):185-196. Fimbria fimbriata:
5(1):21-30 (passim). Gasterosteus
aculeatus: 5(2):185-196. Gastropoda,
Unspecified: 4(1):103. Glaucus atlan-
ticus, Haminoea navicula, Haplo-
chromis burtoni, Hopkinsia rosacea:
5(2):185-196. (Hypselostyla):
3(1):98-99. //vanassa obsoleta:
$1:35-50. Jorunna tormentosa:
5(2):185-196. Laevicaulis alte:
$1:35-50. Laicus argentatus:
5(2):185-196. Lampsilis radiata
luteola: 2:86. Limax maxima: 2:78.
Littorina irrorata: 2:78; S1:35-50. Lot-
tia gigantea: 2:80; S1:35-50. Lym-
naea palustris: S1:35-50. Macoma
balthica: 5(1):21-30 (passim). Mulinia
lateralis: 2:35-40. Musculium securis:
5(1):21-30 (passim). Onchidium ver-
ruculatum: 1:13 (passim). Periploma
spp.: 2:35-40. Petromyzon marinus:
5(1):21-30 (passim). Phestilla spp.,
Phyllaplysia zostericola, Phyllodes-
mium spp. Pinufius rebus:
5(2):185-196. Pisidium spp.:
5(1):21-30. Polymesoda (Geloina)
erosa: 5(1):21-30 (passim), 91-99.
Rossia pacifica: 2:91-92. Rostanga
spp.: 5(2):185-196. Serripes groen-
landicus: 2:94. Siphonaria alternata:
$1:35-50. Sphaerium spp.: 5(1):21-30
(all passim). Spisula solidissima: 1:13
(passim); 2:35-40. Spurilla
neapolitana, Tergipes tergipes:
5(2):185-196. Tritonia: 2:78. T.
diomeda: 1:13 (passim); 2:78. T.
nilsodhneri: 5(2):185-196. Yoldia
hyperborea: 2:94
Behavior, Deimatic
Opisthobranchia: 5(2):185-196
Berry, S. Stillman
Biography, Obituary: 3(1):55-61.
Taxa, Publications: 3(1):63-82
Biochemistry
Amoeba proteus, Biomphalaria
glabrata, Chilomonas, Colpidium,
Crassostrea virginica, Daphnia,
Liolophura gaimardi, Monas, Mya
arenaria, Mytilus edulis, Periplaneta
americana, Schistosoma mansoni:
$1:79-83
Bioenergetics
Australorbis glabratus, Biomphalaria
glabrata, Helisoma trivolvis, Lymnaea
_(Stagnicola) elodes, Lymnaea
palustris, Macoma balthica, Mytilus
edulis, Planorbis corneus:
3(2):213-221

Biofouling
Balanus improvisus: S2:133-142.
Bythinia tentaculata: S2:1-5 (passim).
Corbicula: S2:1-5, 41-45, 47-52, 53-58,
59-61, 63-67, 83-88, 95-98. C. fluminea:
S2:7-39 (passim), 69-81, 99-111, 113-124.
Mytilus: S2:1-5 (passim)

Biofouling Control
Corbicula: S2:41-45, 47-52, 53-58,
59-61, 63-67, 83-88, 95-98. C.
fluminea: S2:69-81

Biological Control
Achatina fulica: 2:98-99. Biomphalaria
spp., Croton sp.-09: 1:67-70. Euglandia
rosea, Gonaxis kibweziensis, Gonaxis
quadrilateralis: 2:98-99. Lymnaea
(Stagnicola) elodes: 1:67-70

Biomass
Corbicula fluminea: 1:96

Biotelemetric Transmitters
Mollusca, unspecified: 1:89

Blood
Melampus bidentatus: 4(1):110-111

Blood Typing, Human
Pulmonata: 1:97-98

Brooding
Acmaeidae: 2:95. Calyptraeidae, Calyp-
traea spp., Crepidula spp., Crucibulum
spp., Hipponix grayanus: 4(2):173-183.
Scaeurgus patagiatus, S. unicirrhus:
6(2):207-211. Transennella tantilla: 2:94

Buoyancy
Nautilus macromphalus: 2:90

Byssus
Anomia simplex: 1:101-102; 2:41-50. Ar-
cacea: 2:41-50. Bivalvia, Unspecified:
4(1):102-103. Boonea impressa: 3(1):97.
Gastropoda, Unspecified: 4(1):102-103.
Mytilacea, Mytilus edulis: 2:41-50.
Odostomia impressa: 3(1):97. Ostreidae:
2:41-50. Pandoracea, Pectinacea:
2:41-50

C-Banding Technique
Ashmunella lenticula, A. proxima
albicaudata: 1:106

Calcite
Anomia simplex: 2:41-50

Calcium, Shell
Ancylus fluviatilis, Biomphalaria
glabrata, B. pfeiffferi, Cincinnatia cincin-
natiensis (passim), Ferrissia rivularis
(passim), Helisoma anceps (passim),
Lymnaea (Stagnicola) elodes, L.
peregra (passim), Nucella lapillus
(passim), Physella gyrina (passim), P. in-
tegra (passim): 5(1):105-124. Pinctada
martensi: 1:101. Planorbis corneus,
Sphaerium spp., Valvata tricarinata:
5(1):105-124 (all passim)

Canadian National Mollusc Collection
New quarters: 2:81

Carbohydrates
Cionella lubrica: 3(1):27-32. Octopus

dolfleini: 2:91
Celestial Cues
Aplysia brasiliana: 2:78
Chemoreceptive Structures
Achatina fulica, Aplysia californica:
2:78
Chromata, absence of
Crassostrea: 1:35-42. Ostrea: 1:90
Chromosomes
Biomphalaria glabrata, B. straminea,
Bulinus tropicus: 1:106-107
Cilia
Corbicula fluminea: 1:13-20. Nucula
sulcata: 1:16 (passim)
Circulatory System
Cerithidea scalariformis: 2:1-20
Cladistic Analysis
Boretrophon aculeatus: 3(1):11-26.
Cerithidea, Cerithideopsilla, Cerithi-
deopsis: 2:1-20. Nucella lamellosa,
Paziella pazi, Trophon geversianus:
3(1):11-26
Climate
Pelecypoda, Unspecified: 2:79
Color
Collisella pelta: 2:80. Cyphoma gib-
bosus: 2:84
Color Patterns, Body
Monadenia, M. fidelis: 3(1):3 (passim)
Competition
Littorina littorea, L. obtusata: 1:92
Condition Index
Polymesoda caroliniana: 6(2):199-206
Conditioning
Lymnaea stagnalis: 2:78
Cooper, James Graham
Biography: 1:89
Copper Toxicity
Pomacea paludosa, Stagnicola sp.:
1:97
Cryptic Coloration
Opisthobranchia: 5(2):185-196,
243-258
Crystalline Style
Ilyanassa obsoleta: 4(1):110
Ctenidium
Adula falcata (passim), Arcuatula:
5(2):159-164. Bankivia, Gastropoda,
Unspecified: 3(1):95. Limnoperna,
Modiolus, Musculista: 5(2):159-164
(passim). Perna viridis: 4(2):233;
5(2):159-164. Trochidae, Turritellidae,
Umbonium, Vermatidae: 3(1):95
Cuttlebone
Sepia officinalis, S. orbignyana: 2:91
Dahllite
Lithophaga nigra, Pinctada martensi:
1:101
Deep-Sea
Amygdalum, Modiolus, Musculus,
Myrina: S1:23-34
Degrowth
Adalaria proxima: 4(1):103-104.

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 289

Australorbis glabratus, Biomphalaria
glabrata: 3(2):213-221. Helisoma
anceps: 4(1):118-119. H. trivolvus:
3(2):213-221; 4(1):118-119. Lymnaea
(Stagnicola) elodes, Macoma
balthica, Mytilus edulis: 3(2):213-221.
Onchidoris muricata: 4(1):103-104.
Planorbis corneus, Polycelis tenuis
(passim), Scrobicularia (passim),
Tellina (passim): 3(2):213-221
Development
Acanthodoris spp.: 5(2):197-214. Ac-
maeidae, Acochlidiacea: 2:95. Acteo-
cina canaliculata, Acteonia cocksi,
Adalaria: 5(2):197-214. A. proxima:
4(1):103-104; 5(2):197-214. Aegires
spp.: 5(2):197-214. Aeolidiella alderi,
A. sanguinea: 5(2):303-306. Aglaja
ocelligera, Aldaria modesta, Aldisa
spp.: 5(2):197-214. Amnicola winkleyi:
4(1):101-102. Ancula pacifica,
Anisodoris nobilis, Antonietta
luteorufa, Aplysia juliana, Aplysiopsis
smithi, Archidoris odhneri:
5(2):197-214. A. pseudoargus:
4(1):103-104; 5(2):197-214. Argonauta
argo: 5(2):303-306. Armina califor-
nica, A. maculata: 5(2):197-214.
Astraea rugosa: 5(2):303-306.
Australorbis glabratus: 3(2):213-221.
Babaina: 5(2):197-214. Berthelinia
caribbea, Berthella californica, Berth-
ellina citrina: 5(2):197-214. Biom-
phalaria glabrata: 3(2):213-221.
Bosellia mimetica: 5(2):197-214.
Cadlina laevis: 4(1):103-104;
5(2):197-214. Cadlina modesta,
Caliphylla mediterranea, Calliopaea
bellula, Calma glaucoides, Calmella
carolinii: 5(2):197-214. Calyptogena
magnifica: 4(1):49-54. Casella ob-
soleta, Catriona gymnota, C. maua,
Chelidonura, Chromodoris spp.:
5(2):197-214. Cincinnatia winkleyi:
4(1):101-102. Corbicula fluminea:
2:87; 4(1):61-79, 81-88, 115-116;
$2:69-81. Costasiella ocellifera:
5(2):197-214. Crassostrea virginica:
$3:41-49, 59-70. Cratena peregrina,
Crimora coneja, C. papillata:
§(2):197-214. Cryptomphalis (Helix)
aspersa: 5(2):303-306. Cryptozona
belangeri: 4(2):237. Cumanotus
beaumonti, Cuthona spp., Cyerce
cristallina: 5(2):197-214. Cylinchnella
canaliculata: 1:91. Dendrodoris spp.,
Dendronotus spp., Dermatobranchus
Striatellus, Diaphana californica,
Dicata odhneri, Dirona albolineata,
Dirona aurantia, Discodoris spp.,
Doridella obscura, D. steinbergae,
Doriopsilla pharpa, Doris, D.
ocelligera, Doto spp., Elysia spp.:

5(2):197-214. Embletonia pulchra:
5(2):303-306. E. pulchra faurei,
Eolidina mannarensis: 5(2):197-214.
Epitonium albidum: 1:1-12. Ercolania
funerea, E. fuscata, Eubranchus
spp., Facelina spp., Fiona pinnata,
Flabella spp., F. affinis: 5(2):197-214.
Gastropoda, Unspecified: 4(1):103.
Glossodoris spp., Goniodoris
castanea, Gymnodoris striata,
Hallaxa chani, Haminoea spp., Han-
cockia ucinata: 5(2):197-214.
Hedylopsis spiculifera: 5(2):303-306.
Helisoma trivolvis: 3(2):213-221. Her-
maea bifida, Hoplodoris nodulosa:
5(2):197-214. Hydrobia truncata:
4(1):101-102. Hypselodoris bennetti,
H. messinensis: 5(2):197-214. Illex il-
lecebrosus: 2:51-56; 4(1):55-60.
Jorunna tormentosa: 5(2):185-196.
Lalia cockerelli, Limapontia capitata,
Limenanara nodosa: 5(2):197-214.
Lissarca notocadensis: 4(2):235.
Lobiger serradifalci: 5(2):197-214.
Lymnaea (Stagnicola) elodes, L.
palustris, Macoma balthica:
3(2):213-221. Melanochlamys
diomedea, Melibe fimbriata, M.
leonina, Miamira sinuata:

5(2):197-214. Moroteuthis pacifica, M.

robusta: 4(2):241. Mytilus edulis:
3(2):213-221. Nucella emarginata:
1:105. N. lapillus: 4(1):110. Octopus
burryi: 2:92. O. dofleini martini:
4(2):241. Oenopopta fidicula,
Oenopota levidensis: 2:94-95.
Okadaia elegans, Olea hansineensis,
Onchidoris bilamellata: 5(2):197-214.
O. muricata: 4(1):103-104;
5(2):197-214. O. neapolitana, Oxynoe
azuropunctata, Peltodoris
atromaculata, Phestilla melano-
branchia, P. sibogae, Phidiana
crassicornis, Philine gibba,
Phyllaplysia engeli, P. taylori,
Phylliroe bucephala, Piseinotecus
sphaeriferus: 5(2):197-214. Pisidium
casertanum: 4(1):116. Placida cremo-
niana, P. viridis: 5(2):197-214. Planor-
bis corneus: 3(2):213-221. Platydoris
scabra, Polycera quadrilineata, P
zosterae, Polycerella emertoni:
5(2):197-214. Pontohedyle milasche-
witschii: 5(2):303-306. Precuthona
divae: 5(2):197-214. Pseudovermis:
2:95. Pteraeolidia ianthina, Retusa
obtusa: 5(2):197-214. Rissoa parva:
5(2):303-306. Rostanga pulchra,
Runcina ferruginea, R. setoensis,
Scyllaea pelagica, Sebradoris cross-
landi: 5(2):197-214. Solemya reidi:
2:94. Sphaerium striatinum: 4(1):116.
Spurwinkia salsa: 4(1):101-102.

Diet

Digestion

Dispersal

Divergence

Diversity

Dredging

Ecogenetics

Stiliger fuscovittatus, Tenellia pallida,
Tergipes tergipes, Tethys fimbria:
5(2):197-214. Thais emarginata: 1:105.
T. haemastoma canaliculata:
6(2):189-197. Thecacera pennifera,
Thordisa filix, Thorunna spp.,
Trapania maculata, Tridachia crispata,
Triopha catalinae, Trippa spongiosa,
Tritonia diomeda, T. festiva:
5(2):197-214. I. hombergi:
4(1):103-104; 5(2):197-214. Tritoniopsis
cincta: 5(2):197-214. Unela glanduli-
fera: 5(2):303-306. Viviparus
georgianus: 3(2):268

Cyphoma gibbosus: 2:84.
Glossiphona complanata: 5(1):73-84.
llyanassa obsoleta: 4(1):110. Lymnaea
peregra, Planorbis vortex (passim):
5(1):73-84

Cardiomya planetica: 1:13 (passim)

Corbicula: S2:1-5. C. fluminea:
$2:7-29, 231-239

Nucella emarginata, Thais
emarginata: 1:105

Crassostrea spp.: 1:108

Crassostrea virginica: S3:1-4, 5-10, |
11-16, 37-40. Ostrea chilensis: S3:1-4

Theba pisana: 1:104

Ecology

Abra alba: 5(1):21-30 (passim).
Acanthophora spicifera: 5(2):259-280
(passim). Aciculidae: 3(2):223-231.
Acochlidiacea: 2:95; 5(2):281-286.
Acropora palmata: 1:1-12. Actinonaias
ellipsiformis: 3(1):93. A. pectorosa:
3(1):104. Adalaria proxima:
4(1):103-104; 4(2):235; 5(2):293-301.
Adipicola: S1:23-34. Aeolidiella alderi,
A. sanguinea: 5(2):303-306.
Alasmidonta minor: 3(1):104. A.
viridis: 5(1):1-7. Alvania auberiana:
4(2):185-199. Amnicola limosa:
3(1):99; 5(1):9-19, 31-39, 73-84. A.
winkleyi: 4(1):101-102. Ampullariidae:
3(2):223-231. Amygdalum, A.
politum: S1:23-34. Ancylus fluviatilis:
3(2):135-142, 151-168, 243-265;
5(1):105-124. Ankylastrum capuloides,
A. fluviatile: 5(1):65-72 (passim).
Anodonta spp.: 3(1):47-53, 93;
4(2):230-231; 5(1):1-7, 31-39, 41-48,
91-99; 6(2):165-178; S2:1-5. Anodon-
toides ferussacianus: 3(1):93. Antho-
pleura elegantissima, Antiopella bar-
barensis: 5(2):287-292. Aplacophora:
3(1):93-94; 5(2):281-286; S1:23-34.

290 AMER

Aplysiopsis zebra: 5(2):259-280.
Archaeogastropoda: $1:23-34. Arch-
idoris pseudoargus: 4(1):103-104.
Arctica islandica: S3:51-57.
Arenicola: 2:96. Argonauta argo:
5(2):303-306. Ascobulla ulla:
5(2):259-280. Aspidodiadema
hawailiensis: 2:83. Assimineidae:
3(2):223-231. Astarte castanea:
5(1):21-30 (passim). Astraea rugosa:
5(2):303-306. Atrina seminuda: 2:97.
Australorbis glabratus: 3(2):213-221.
Bankia gouldi: 4(1):89-99;
$1:101-109. Batissa (Cyrenobatissa)
subsulcata: 5(1):91-99. Berthelinia
caribbea: 5(2):259-280. Biom-
phalaria spp.: 3(2):213-221; 4(1):120;
5(1):65-72, 85-90. Bithynia:
3(2):135-142 (passim), 269-272.
Bithyniidae: 3(2):223-231. Bivalvia,
Unspecified: 3(1):93-94;
4(1):102-103; 6(1):49-54. Bosellia
mimetica, Bosellidae: 5(2):259-280.
Brechites penis: 5(1):21-30 (passim).
Buccinum undatum: 3(2):223-231.
Bulinus jousseaumei: 5(1):65-72. B.
truncatus: 5(1):85-90. Bythinia ten-
taculata: 5(1):65-72 (passim).
Cadlina laevis: 4(1):103-104.
Caecum: 5(2):281-286. C. nitidum:
4(1):185-199. Caliphyllidae:
5(2):259-280. Callinectes sapidus:
$3:51 (passim). Calyptogena:
$1:23-34. C. magnifica: 4(1):49-54;
$1:23-34. C. ponderosa: $1:23-34.
Calyptraeidae: 4(2):173-183. Calyp-
traea spp.: 4(2):173-183. Campeloma
decisum: 5(1):9-19, 31-39, 73-84,
101-104. Catriona gymnota:
5(2):287-292. Caulerpa mexicana, C.
sertulariodes: 5(2):259-280. Cepaea
nemoralis: 5(1):105-124. Cerithidea
spp.: 2:1-20. Cerithiidae:
3(2):223-231. Chaetomorpha:
5(2):259-280. Chondrocidaris
gigantea: 2:83. Chromodoris, C.
albopunctatus, Cimora coneja:
5(2):287-292. Cincinnatia cincinna-
tiensis: 5(1):31-39. C. winkleyi:
4(1):101-102. Cipangopaludina
chinensis: 5(1):9-19. Cladophora
gracilis: 5(2):259-280 (passim).
Codakia orbicularis: S$1:23-34.
Codium isthmocladium:
5(2):259-280. Corbicula: 5(1):21-30
(passim); S2:41-45, 47-52, 53-58,
63-67, 83-88, 95-98. C. fluminalis:
5(1):91-99; S2:203-209. C. fluminea:
1:96; 3(1):41-45, 94; 3(1):100,
100-101; 3(2):267-268; 4(1):61-79;
5(1):1-7, 31-39, 91-99; S2:7-39, 69-81,
89-94, 99-111, 133-142, 143-150,
151-166, 167-178, 179-184, 203-209,

211-218, 219-222, 223-229, 231-239.
C. leana: 4(1):81-88; S2:202-209.
Corbiculacea: 3(2):201-212;
5(1):21-30 (passim). Cordylophora
lacustris, Coryphella spp.:
5(2):287-292. Costasiella ocellifera,
C. nonatoi, Costasiellidae:
5(2):259-280. Crassostrea virginica:
$1:111-116; S3:1-4, 5-10, 25-29, 31-36,
41-49, 59-70, 71-75. Crenella:
$1:23-34. Crepidula convexa:
3(1):33-40; 4(2):173-183. C. fornicata:
3(2):135-142 (passim); S2:203-209. C.
plana: 3(1):33-40; 4(2):173-183. C.
spp. Cristaria (Pletholophus)
discoidea: 5(1):91-99 (passim).
Crossaster papposis: 5(2):287-292.
Crucibulum spp.: 4(2):173-183. Cryp-
tomphalis (Helix) aspersa:
5(2):303-306. Cuthona spp.:
5(2):287-292. Cyclonaias tuberculata:
2:85. Cyclophoridae: 3(2):223-231.
Cyerce antillensis: 5(2):259-280.
Dacrydium: S1:23-34. Dendronotus
diversicolor: 5(2):287-292. Deroceras
reticulatum: 3(2):223-231. Donax
fossor: 3(1):92. Dreissena poly-
morpha: 5(1):91-99 (passim). Elliptio
cistelliformis: 1:61-68. E. complanata:
5(1):31-39. E. crassidens: 3(1):41-45;
6(2):165-178. E. crassidens
crassidens: 4(1):117. E. dilatata:
3(1):41-45; 6(2):165-178. E. spp.:
1:61-68. Elysia: 5(2):287-292. E. spp.,
Elysiidae: 5(2):259-280. Embletonia
pulchra: 5(2):303-306. Enis: 2:96.
Epioblasma capsaeformis:
6(2):165-178. Epitonium albidum:
1:1-12. Ercolania funerea, E. fuscata:
5(2):259-280. Eubranchus:
5(2):243-258. E. sanjuanensis, E.
tricolor: 5(2):287-292. Eupera cuben-
SiS: $2:223-229. Facelina bostonien-
Sis: 5(2):287-292. Falcidens:
$1:23-34. Ferrissia: 5(1):73-84. F.
fragilis: 3(1):99; 5(1):9-19. F. paraliela:
5(1):9-19. F. rivularis: 3(2):135-142
(passim). Fimbria fimbriata:
5(1):21-30 (passim). Fossaria
modicella: 3(1):99. Fusconaia barne-
Siana: 3(1):41-45, 104; 6(2):165-178. F.
barnesiana bigbyensis: 3(1):41-45;
5(1):1-7. F ebena: 5(2):177-179. F.
edgariana: 3(1):104. F. flava: 3(1):93.
F. ozarkensis: 2:85. F. subrotunda:
3(1):41-45. Gafrarium pectinatum:
5(1):91-99 (passim). Gastrohedyle:
5(2):281-286. Gastropoda,
Unspecified: 3(1):93-94; 4(1):102-103,
114; 5(1):101-104. Gemma gemma:
2:96. Geukensia demissa demissa:
5(1):173-176. Geukensia demissa
granosissima: 3(1):103; 4(1):112;

. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

5(1):173-176. Glycera: 2:96. Granulina
ovaliformis: 4(1):185-199. Gyraulus cir-
cumstriatus, G. deflectus: 3(1):99;
5(1):9-19. G. parvus: 5(1):9-19, 31-39,
73-84. Haliotis cracherodii:
4(2):233-234. H. corrugata:
3(2):223-231. H. roei: 3(1):97. H.
rufescens: 3(2):223-231. Hancockia
californica: 5(2):287-292. Hedylopsis:
5(2):281-286. H. spiculifera:
5(2):303-306. Helicinidae:
3(2):223-231. Helisoma anceps:
3(1):99; 5(1):9-19, 31-39, 73-84. H.
campanulatum: 3(1):99; 5(1):9-19. H.
trivolvis: 3(2):213-221; 5(1):9-19. H.
pomatia: 3(2):223-231. Hermissenda
crassicornis: 5(2):287-292. Hiatella:
3(2):135-142 (passim). Hipponix
grayanus: 4(2):173-183. Hydrobia
truncata: 4(1):101-102. Hydrobiidae,
Hydrocenidae: 3(2):223-231. /dasola,
I. argentea: S1:23-34. Illex il-
lecebrosus: 4(1):55-50, 101; 4(2):239.
Ilyanassa obsoleta: 2:14 (passim).
Jorunna tormentosa: 4(1):103-104.
Lacuna cossmanni: $1:23-34.
Lacuna vincta: 5(2):287-292.
Laevapex fuscus: 3(1):99; 5(1):9-19.
Lalia cockerelli: 5(2):287-292.
Lamellibranchia: S1:23-34. Lamprotula
leai: 5(1):91-99. Lampsilis spp.:
1:61-68; 2:85; 3(1):41-45, 93, 104;
4(2):230-231; 5(1):1-7, 31-39;
6(2):165-178. Lasmigona compressa:
3(1):93. L. costata: 3(1):104;
6(2):165-178. Leptodea fragilis:
6(2):165-178. Leptosynapta: 2:96.
Leptoxis carinata: 3(2):169-177;
4(1):119. Lexingtonia dolabelloides:
3(1):104. Ligumia subrostrata:
5(1):41-48. Limapontia capitata:
5(2):259-280. Limax pseudoflavus:
3(2):223-231. Limnoperna fortuei:
5(1):91-99. L. lacustris, L. supoti:
5(1):91-99 (passim). Littorina filosa:
4(1):112. L. irrorata: 3(2):223-231. L.
littorea: 3(2):135-142 (passim). L.
mespillium: 4(1):185-199. L. saxatilis:
1:92-93. L. scabra: 4(1):112. Lobiger |
souverbiei: 5(2):259-280. Loligo
opalescens: 4(1):55-50; 4(2):240. L.

peali: 4(1):101. L. vulgaris: 4(1):55-50.
Lottia gigantea: 2:80; 4(2):242-243.
Lucina atlantis, L. (Linga) penn-
sylvanica, L. (Phacoides) pectinatus,
Lucinidae, Lucinoma, L. atlantis, L.
filosa: $1:23-34. Lymnaea

(Stagnicola) elodes: 3(2):143-150,
213-221; 5(1):73-84, 105-124 (passim);
6(1):9-17. L. emarginata: 5(1):73-84.
L. palustris: 3(2):213-221. L. peregra:
3(2):135-142 (passim); 5(1):65-72,
73-84. L. stagnalis: 3(2):135-142

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 291

(passim), 223-231; 5(1):65-72.
Lyogyrus granum: 5(1):9-19. Lyonsia
californica: 5(1):173-176 (passim).
Lysinoe, L. ghiesbreghti:
3(1):102-103. Macoma balthica:
3(2):213-221; 5(1):21-30 (passim). M.
calcarea: 2:94. Magnonaias nervosa:
4(2):230-231. Maraunibina verrucosa:
5(2):281-286. Margaritifera
margaritifera: 5(1):91-99 (passim);
§(2):125-128. Marisa cornuarietis:
3(2):223-231. Mazatlania aciculata:
1:92. Medionidus conradicus:
3(1):41-45, 104; 5(1):1-7; 6(2):165-178.
Meiomenia, Meiopriapulus fijiensis:
5(2):281-286. Melampus bidentatus:
3(2):135-142 (passim). Melaniidae:
3(2):223-231. Melanoides tuber-
culata: 5(1):105-124. Melanoposidae:
3(2):223-231. Mellanella sp.: 2:83.
Mercenaria: 2:96. Mercuria confusa,
M. punica: 5(1):85-90. Meso-
gastropoda: 3(2):223-231; $1:23-34.
Metridium senile: 5(2):287-292.
Micromenetus dilatatus: 3(1):99;
5(1):9-19. Modiolus: S1:23-34. M.
modiolus: 4(1):104. Mogula:
5(2):287-292 (passim). Mollusca,
Unspecified: 3(1):96-97, 107;
3(2):135-142 (passim). Mourgona ger-
maineae: 5(2):259-280. Mulinia sp.:
4(1):104. Musculium lacustre:
3(2):187-200; 5(1):91-99. M. par-
tumeium: 3(2):187-200, 201-212;
$2:223-229. M. securis: 3(2):187-200;
5(1):21-30 (passim), 31-39; S2:223-229.
M. transverum: S2:223-229.
Musculus: $1:23-34. Mya: 2:96. M.
truncata: 2:94. Myrina: $1:23-34.
Mysella tumida: 4(2):234. Mytilidae:
3(1):95; S1:23-34. Mytilimera nutalli:
5(1):173-176 (passim). Mytilopsis
leucophaeta, M. sallei: 5(1):91-99
(passim). Mytilus: 5(1):41-48. M.
edulis: 3(2):213-221; 4(1):104;
5(1):91-99 (passim). M. galloprovin-
Cialis: 5(1):91-99 (passim). Navanax
inermis: 5(2):287-292. Neogastro-
poda, Neomenia: S1:23-34.
Neomeniomorpha: 5(2):281-286.
Neomphalace, Neomphalidae, Neom-
phalus fretterae: S1:23-34.
Neopisidium: S2:223-229. Nereis:
2:96. Nerita fulgurans, Neritacea,
Neritidae, Neritina latissima:
3(2):223-231. Nudibranchia: 2:84;
5(2):281-286. Obelia: 5(2):287-292
(passim). Obovaria: 4(2):230-231. Oc-
topus bimaculoides: 2:90;
4(2):241-242. O. briareus: 6(1):45-48.
O. dolfleini: 2:90; 6(1):45-48. O.
tetricus, O. vulgaris: 6(1):45-48.
Onchidoris aspersa: 5(2):293-301.

O. bilamellata: 5(2):287-292. O.
muricata: 4(1):103-104; 5(2):293-301.
Opisthobranchia: 5(2):281-286.
Opuntia littoralis: 2:98. Ostrea chilen-
sis: $3:1-4. Oxynoe antillarum, O.
azuropunctata: 5(2):259-280.
Paraganitus ellynnae: 5(2):281-286.
Patella vulgata: 3(2):223-231. Patelli-
dae: 3(1):95. Pelseneeria spp.: 2:83.
Periploma margaritaceum, P. orbicu-
lare: 2:35-40. Perna viridis:
5(2):159-164. Petromyzon marinus:
5(1):21-30 (passim). Phestilla:
5(2):287-292. Philinoglossa, P_ mar-
cusi: 5(2):281-286. Physa ancillaria:
5(1):9-19. P. fontinalis: 3(2):135-142
(passim); 5(1):65-72 (passim). P
heterostropha: 5(1):9-19. P. integra:
5(1):73-84. P. propinqua: 5(1):65-72
(passim). Physella ancellaria: 3(1):99.
P. gyrina: 5(1):31-39. P. virgata
virgata: 3(2):243-265. Pinctada
martensi: 5(1):173-176 (passim). Pin-
nidae: 2:97. Pisidiidae: 3(2):201-212.
Pisidium spp.: 3(2):187-200, 201-212;
5(1):1-7, 21-30, 31-39, 41-48, 49-64,
91-99; S2:223-229. Placida den-
dritica, P kingstoni: 5(2):259-280.
Placopecten magellanicus: 4(1):104;
6(1):1-8. Planorbis corneus:
3(2):135-142 (passim), 213-221. P
planorbis, P. vortex: 5(1):65-72.
Planorbula armigera: 3(1):99;
5(1):9-19. Pleurobema coccineum:
2:85. P. oviforme: 3(1):41-44, 104;
5(1):1-7; 6(2):165-178. Pleurobranchaea
californica: 5(2):287-292.
Pleuroceridae: 3(2):223-231.
Pogonophora: S1:23-34. Polinices
duplicatus: 3(2):135-142 (passim).
Polymesoda caroliniana:
6(2):199-206. P. (Geloina) erosa:
5(1):21-30 (passim), 91-99. Pomacea
lineata: 3(2):223-231. Pontohedyle
milaschewitschii: 5(2):303-306.
Potamilus alatus: 3(1):41-45;
6(2):165-178. P capax: 4(2):230-231.
Potamopyrgus jenkinsii: 3(2):223-231;
5(1):73-84. Prionocidaris hawaiiensis:
2:83. Promenetus exacuous: 3(1):99;
5(1):9-19. Prosobranchia, Pseudo-
miltha: S1:23-34. Pseudopleuronectes
americanus: 5(2):287-292. Pseudo-
succinea columella: 3(1):99; 5(1):9-19.
Pseudovermis: 2:95; 5(2):281-286. P
hancocki, P mortoni, Pseudunela, P
cornuata: 5(2):281-286. Ptycho-
branchus fasciolaris: 3(1):104. P. oc-
cidentalis: 2:85. P subtentum:
3(1):104. Puperita pupa: 4(1):185-199.
Quadrula fragosa, Q. metanerva:
4(2):230-231. Q. pustulosa:
6(2):165-178. Radiocentrum

avalonense: 2:98. Radix limosa:
5(1):65-72 (passim). R. quadrasi:
5(1):105-124 (passim). Rissoa parva:
5(2):303-306. Rissoella caribaea:
4(2):185-199. Rissoidae: 3(2):223-231.
Rissoina bryera, R. catesbyana:
4(2):185-199. Salvia mellifera: 2:98.
Sargassum: 5(2):259-280 (passim).
Scaphopoda: 3(1):93-94. Scoloplos:
2:96. Semibalanus balanoides:
$1:111-116. Sepietta oweniana: 2:90.
Setoaeolis pilata: 5(2):287-292.
Simrothiella, Simrothiellidae:
$1:23-34. Smaragdia viridis viride-
maris: 4(2):185-199. Solemya
(Acharax) spp., S. agassizi: $1:23-34.
S. reidi: 2:94. S. velum, Solemyidae:
$1:23-34. Soletellina elongata: S2:1-5
(passim). Sphaerium spp.:
3(2):187-200, 201-212; 5(1):1-7, 21-30,
31-39, 41-48, 91-99; S2:223-229.
Spirodon carinata: 3(2):169-177.
Spisula solidissima: 3(2):135-142
(passim). Stagnicola elodes:
5(1):9-19. S. palustris: 5(1):65-72
(passim). Stiligeridae: 5(2):259-280.
Strombus gigas: 3(2):223-231.
Strophitus undulatus: 4(1):41-45.
Stylpopodium zonale: 5(2):259-280
(passim). Syllis: 2:29. Syrnolopsidae:
3(2):223-231. Tenellia adspersa:
5(2):287-292. Teredo bartschi:
4(1):89-99; S1:101-109; S2:203-209. T.
furcifera: $1:101-109. T. navalis:
4(1):89-99; S1:101-109. Thalassia
testudinum: 5(2):259-280. Theodoxia
fluviatilis: 5(1):65-72 (passim).
Thiaridae: 3(2):223-231. Thyrasira,
Thyrasiridae: S1:23-34. Toxolasma
lividus: 3(1):41-45, 104; 6(2):165-178.
T. pullus: 1:61-68. Tricolia spp.:
4(2):185-199. Triopha catalinae:
5(2):287-292. Tritonia hombergi:
4(1):103-104. Trochacea: S1:23-34.
Trochidae: 3(1):95. Trochostylifer sp.:
2:83. Turridae: S1:23-34. Unela glan-
dulifera: 5(2):303-306. Union
douglasiae: 5(1):91-99. Unionacea:
3(2):201-212. Unionidae, Unspecified:
1:93-94; 3(1):106; 4(1):101; S2:1-5.
Urosalpinx cinerea: S1:111-116.
Valvata tricarinata: 5(1):9-19, 31-39.
Valvatacea, Valvatidae: 3(2):223-231.
Vaucheria: 5(2):259-280. Vesicomya,
V. caudata, V. cordata: S1:23-34.
Vesicomyidae: 3(1):95-96; S1:23-34.
Vestimentifera: S1:23-34. Villosa iris:
3(1):41-45; 6(2):165-178. V. iris iris:
2:85. V. nebulosa: 3(1):104; 5(1):1-7.
V. ogeecheensis: 1:61-68. V. vanuxe-
mensis: 3(1):41-45; 6(2):165-178. V.
vanuxemi: 3(1):104; 5(1):1-7. Vitreolina
sp.: 2:83. Viviparacea, Viviparidae,

292 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Viviparus bengalensis: 3(2):223-231.
V. georgianus: 3(2):268; 5(1):9-19.
V. melleatus, V. viviparous: 3(2):223-231.
Volvatella bermudae: 5(2):259-280.
Yoldia hyperborea: 2:94. Zebina
browniana: 4(2):185-199

Ecology, Chemical
Balanus amphitrite amphitrite:
$1:111-116. Crassostrea virginica:
4(1):101; S1:111-116. Semibalanus
balanoides, Urosalpinx cinerea:
$1:111-116

Ecology, Population
Cepaea spp.: 1:107-108

Egg Capsules
Acteonia cocksi: 4(2):205-216
(passim). Adelomelon brasiliana:
4(2):165-172. Aeolidacea (passim),
Aeolidia papillosa: 4(2):205-216.
Alloteuthis: 4(2):217-227. Alvania
spp.: 4(1):185-199. Aplysia punctata,
Archidoris spp.: 4(2):205-216
(passim). Argonauta: 4(2):217-227.
Armina tigrina: 4(2):205-216 (passim).
Assiminea californica: 4(1):185-199
(passim). Austrodoris macmurdensis:
4(2):205-216 (passim). Bathypolypus
arcticus: 4(2):217-227. Buccinum un-
datum, Busycon sp., B. carica:
4(1):185-199 (passim). Cadlina laevis:
4(2):205-216 (passim). Caecum
nitidum, Calliostoma zizyphinum
(passim), Calotrophon ostrearum
(passim): 4(1):185-199. Calyptraeide:
4(2):173-183. Calyptraea spp.:
4(2):173-183. Cantharus multangulus:
4(1):185-199 (passim). Cerithidea
californica: 4(2):165-172. Cingula:
4(1):185-199 (passim). Conus:
4(2):229. C. figulinus, C. jaspideus
stearnsi: 4(1):185-199 (passim). Cory-
phella salmonacea, Costasiella
lilanae: 4(2):205-216. Crepidula for-
nicata: 4(2):165-172. C. spp.,
Crucibulum spp.: 4(2):173-183. Den-
drodoris albopunctata, Dendronotus
frondosus: 4(2):205-216. Eledone
cirrhosa, E. moschata, Eledonella
pygmaea: 4(2):217-227. Elysia cauze,
Embletonia fuscata: 4(2):205-216
(passim). Epitonium albidum: 1:1-12;
4(1):185-199 (all passim). Eupleura
caudata: 4(1):185-199 (passim).
Euprymna: 4(2):217-227. Granulina
ovaliformis: 4(1):185-199. Haminoea
vesicula: 4(2):165-172. Hermissenda
crassicornis: 4(2):205-216. Hipponix
grayanus: 4(2):173-183. Hyalina
avena: 4(1):185-199 (passim).

. Idiosepius, Illex: 4(2):217-227. Il-
yanassa obsoleta: 4(2):165-172.
Lamellaria perspicua (passim), Lit-
torina mespillium: 4(1):185-199. Loligo

Egg

vulgaris: 4(2):217-227. Marginella
aureocincta, Melarpha cincta
(passim), Murex fulvescens (passim):
4(1):185-199. Nassarius obsoleta, N.
trivittatus: 4(2):165-172. Nautilus:
4(2):217-227. Nerita spp., Neritina
virginea, Nitesselata: 4(1):185-199 (all
passim). Nucella /apillus:
4(2):165-172. Octopodidae, Octopus
spp.: 4(2):217-227. Onoba:
4(1):185-199 (passim). Phyllaplysia
taylori: 4(2):205-216 (passim).
Polinices sp., Polystira barrettii:
4(1):185-199 (passim). Pteroctopus
tetracirrhus: 4(2):217-227. Puperita
puap, Rissoa albella (passim),
Rissoella caribaea, Rissoina bryerea,
R. catesbyana: 4(2):185-199. Rossia,
Sepia, S. elegans: 4(2):217-227. S.
Officinalis: 4(2):165-172, 217-227. S.
orbignyana, Sepietta, Sepiola:
4(2):217-227. Smaragdia viridis
viridemaris: 4(2):185-199. Spirula:
4(2):217-227. Strombus: 4(1):185-199
(passim). Tegula pfeifferi:
4(2):165-172. Tenellia pallida:
4(2):205-216 (passim). Thais
haemastoma canaliculata:
6(2):189-197. T. lapillus: 4(2):165-172.
Theodoxus fluviatilis: 4(1):185-199
(passim). Tremoctopus: 4(2):217-227.
Tricolia spp.: 4(2):185-199. Tritonia
hombergi: 4(2):205-216 (passim),
Urosalpinx cinerea: 4(2):165-172. U.
perrugata: 4(1):185-199 (passim).
Vampyroteuthis, V. infernalis:
4(2):217-227. Zebina browniana:
4(2):185-199

Laying

Epitonium ulu: 1:10. Thais
haemastoma canaliculata:
6(2):189-197

Eggs

Acanthodoris spp., Acteocina
canaliculata, Acteonia cocksi,
Adalaria, A. proxima: 5(2):197-214.
Adelomelon brasiliana: 4(2):165-172.
Aegires spp., Aglaja ocelligera,
Aldaria modesta, Aldisa spp.:
5(2):197-214. Anaspidea: 4(1):109-110.
Ancula pacifica, Anisodoris nobilis,
Antonietta luteorufa, Aplysia juliana,
Aplysiopsis smithi, Archidoris odhneri,
A. pseudoargus, Armina californica,
A. maculata, Babaina, Berthelinia
caribbea, Berthelinia limax, Berthella
californica, Berthellina citrina,
Bosellia mimetica, Cadlina laevis, C.
modesta, Caliphylla mediterranea,
Calliopaea bellula, Calma glaucoides,
Calmella carolinii: 5(2):197-214.
Calyptraeidae: 4(2):173-183. Casella
obsoleta, Catriona gymnota,

C. maua: 5(2):197-214. Cerithidea

californica: 4(2):165-172. Chelidonura:

5(2):197-214. Chicoreus virgineus:
4(1):109-110. Chromodoris spp.:
4(1):109-110; 5(2):197-214. Conidae:
4(1):109-111. Conus: 4(1):109-110.
Costasiella ocellifera: 5(2):197-214.
Crassostrea virginica: S3:41-49.
Cratena peregrina: 5(2):197-214.
Crepidula spp.: 4(2):165-172, 173-183.
Crimora coneja, C. papillata:
5(2):197-214. Crucibulum spp.:
4(2):173-183. Cumanotus beaumonti,
Cuthona spp., Cyerce cristallina,
Dendrodoris spp., Dendronotus, Der-
matobranchus Striatellus, Diaphana
californica, Dicata odhneri, Dirona
albolineata, D. aurantia, Discodoris
spp., Doridella obscura, D.
steinbergae, Doriopsilla pharpa,
Doris ocelligera, Doto spp., Elysia
spp.: 5(2):197-214. E. olivaceus:
4(1):109-111. Embletonia pulchra
faurei, Eolidina mannarensis, Er-
colania funerea, E. fuscata,
Eubranchus spp., Facelina spp.:
5(2):197-214. Fasciolariidae:
4(1):109-110. Fiona pinnata, Flabella
spp., Flabellina affinis, Glossodoris
spp., Goniodoris castanea:
5(2):197-214. Gymnodoris limaci-
formis: 4(1):109-110. G. striata,
Hallaxa spp.: 5(2):197-214. Haminoea
vesicula: 4(2):165-172; 5(2):197-214.
Hancockia ucinata, Hermaea bifida:
5(2):197-214. Hipponix grayanus:
4(2):173-183. Hoplodoris nodulosa,
Hypselodoris bennetti, H. messinen-
Sis: 5(2):197-214. llyanassa obsoleta:
4(2):165-172. Lalia cockerelli,
Limapontia capitata, Limenandra
nodosa, Lobiger serradifalci, Melano-
chlamys diomedea, Melibe fimbriata,
M. leonina, Miamira sinuata:
5(2):197-214. Murex ramosus,
Muricidae: 4(1):109-110. Nassarius
obsoleta, N. trivittatus: 4(2):165-172.

Nerita forskali, Neritidae: 4(1):109-110.

Nucella lapillus: 4(1):110;
4(2):165-172. Oenopopota fidicula,
Oenopota levidensis: 2:94-95.
Okadaia elegans, Olea hansineenis:
5(2):197-214. Onchidoris aspersa:
5(2):293-301. O. bilamellata:
5(2):197-214. O. muricata:
5(2):197-214, 293-301. O. neapolitana,
Oxynoe azuropunctata, Peltodoris
atromaculata, Phestilla melano-
branchia, P. sibogae, Phidiana
crassicornis, Philine gibba, Phyll-
aplysia engeli, P. taylori: 5(2):197-214.
Phyllida varicosa: 4(1):109-110.
Phylliroe bucephala: 5(2):197-214.

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 293

Phyllobranchillus orientalis, Phyllo-
desmium xeniae: 4(1):109-111.
Piseinotecus sphaeriferus, Placida
cremoniana, P. viridis, Platydoris
scabra: 5(2):197-214. Pleuroploca
trapezuim (sic): 4(1):109-110. Polycera
quadrilineata, P zosterae, Polycerella
emertoni, Precuthona divae,
Pteraeolidia ianthina, Retusa obtusa,
Rostanga pulchra, Runcina fer-
ruginea, R. setoensis: 5(2):197-214.
Sacoglossa: 4(1):109-110. Scy/laea
pelagica: 5(2):197-214. Searlesia dira:
4(2):173-183 (passim). Sebradoris
crosslandi: 5(2):197-214. Sepia of-
ficinalis: 4(2):165-172. Stiliger
fuscovittatus: 5(2):197-214. Strom-
bidae: 4(1):109-110. Tegula pfeifferi:
4(2):165-172. Tenellia pallida,
Tergipes tergipes, Tethys fimbria:
5(2):197-214. Thaididae: 4(1):109-110.
T. lapillus: 4(2):165-172. T.
haemastoma canaliculata:
6(2):189-197. T. savignyi: 4(1):109-110.
Thecacera pennifera, Thordisa filix,
Thorunna spp., Trapania maculata,
Tridachia crispata, Triopha catalinae,
Trippa spongiosa, Tritonia spp..,
Tritoniopsis cincta: 5(2):197-214.
Trochus erythraeus, Turbinidae, Turbo
radiatus: 4(1):109-110. Urosalpinx
cinerea: 4(2):165-172

Eggs, Nurse

Searlesia dira: 4(2):173-183 (passim).
Thais haemastoma canaliculata:
6(2):189-197

Embryology

Corbicula fluminea: 4(1):81-88, 116.
Pisidium casertanum, Sphaerium
Sstriatinum: 4(1):116. Thais haemastoma
canaliculata: 6(2):189-197. Transen-
nella tantilla: 2:94

Endangered Species

Amblema plicata, Dromus dromas:
4(1):117. Dysnomia sulcata delicata,
D. torulosa rangiana, D. triquetra:
3(1):105. Elliptio (Canthyria) stein-
stansana: 3(1):104-104. E. crassidens
crassidens, Epioblasma flexuosa,
Fusconaia subrotunda: 4(1):117.
Lampsilis higginsi: 4(2):230. L. or-
biculata: 2:85, 85-86. L. teres teres,
Ligumia recta, Megalonaias nervosa,
Pleurobema plenum, Potamilus
alatus: 4(1):117. Simpsoniconcha am-
bigua, Villosa fabalis: 3(1):105

Energetics

Corbicula fluminea: S2:143-150.
Onchidoris aspersa, O. bilamellata,
O. muricata: 5(2):293-301

Evolution

Acmaeidae: 4(1):115. Acochlidiacea:
5(2):281-286. Amplirhagada: 1:98-99.

Anomalodesmata: 4(1):111-112.
Aplacophora 5(2):281-286; 6(1):57-68.
Aplysiopsis zebra, Ascobulla ulla:
5(2):259-280. Australorbis glabratus:
3(2):213-221. Bellamya spp.: 4(1):107.
Berthelinia caribbea: 5(2):259-280.
Biomphalaria glabrata: 3(2):213-221.
Bivalvia, Unspecified: 4(1):111-112.
Bosellia mimetica, Bosellidae:
5(2):259-280. Caecum: 5(2):281-286.
Caelatura: 4(1):107. Caliphyllidae:
5(2):259-280. Cellana: 4(1):115.
Chaetomorpha: 5(2):259-280. Con-
voluta convoluta: $1:35-50. Corbicula
fluminea: $1:35-50; S2:223-229.
Costasiella ocellifera, C. nonatoi,
Costasiellidae: 5(2):259-280.
Ctenodonta nasuta: 4(1):111-112.
Cyerce antillensis: 5(2):259-280.
Dacrydium: 4(1):111-112. Elysia spp.,
Elysiidae, Ercolania funerea, E.
fuscata: 5(2):259-280. Eupera cuben-
sis: S2:223-229. Gastrohedyle:
5(2):281-286. Gastropoda,
Unspecified: 2:80-81; 4(2):244.
Hedylopsis: 5(2):281-286. Helisoma
trivolvis: 3(2):213-221. Helix aspersa:
$1:35-50. Heterodonta: 4(1):111-112.
Illex spp.: S1:93-100. Lampsilis:
$1:35-50. Lepetidae: 4(1):115.
Limapontia capitata: 5(2):259-280.
Littorina obtusata: 4(1):108. Lobiger
souverbiei: 5(2):259-280. Loligo:
$1:93-100. Lymnaea (Stagnicola)
elodes, L. palustris, Macoma
balthica: 3(2):213-221. Malletiidae:
4(1):111-112. Maraunibina verrucosa,
Meiomenia, Meiopriapulus fijiensis:
5(2):281-286. Micrarionta opuntia, M.
sodalis: 4(2):237. Mourgona ger-
maineae: 5(2):259-280. Musculium
spp.: S2:223-229. Mytilus edulis:
3(2):213-221. Nautilus macrom-
phalus: S1:93-100. Neomeniomorpha:
5(2):281-286. Neopisidium:
$2:223-229. Neothauma tanganyi-
cense: 4(1):107. Nucinellidae, Nucul-
acea, Nuculanacea: 4(1):111-112.
Nudibranchia: 5(2):281-286. Octo-
podidae, Octopus vulgaris:
$1:93-100. Opisthobranchia:
5(2):281-286. Oxynoe antillarum, O.
azuropunctata: 5(2):259-280. Paleo-
heterodonta: 4(1):111-112. Paraganitus
ellynnae: 5(2):281-286. Patella,
Patellidae, Patellogastropoda:
4(1):115. Pectinacea: 4(1):111-112.
Philinoglossa, P. marcusi:
5(2):281-286. Pisidium spp.:
$2:223-229, Placida dendritica, P
kingstoni: 5(2):259-280. Planorbis
corneus: 3(2):213-221. Pliodon ovata,
P. spekii: 4(1):107. Polyplacophora:

6(1):57-68. Protobranchia:
4(1):111-112. Pseudovermis spp.,
Pseudunela, P. cornuta: 5(2):281-286.
Solemyidae, Solemyoidae: 4(1):111-112.
Sphaerium spp.: S2:223-229.
Stiligeridae: 5(2):259-280. Viviparidae:
3(1):107. Volvatella bermudae:
5(2):259-280. Westraltrachia: 1:98-99

Evolution, Chromosome

Biomphalaria glabrata, B. straminea,
Bulinus tropicus: 1:106-107

Extinction

Ammonites: 2:79. Epioblasma samp-
soni: 1:27-30. Pelecypoda, Unspeci-
fied: 2:79

Eyes

Cephalopoda, Unspecified: 2:90-91.
Cerithidea scalariformis: 4(1):111;
4(2):234. Gourmya gourmyi: 2:1-20.
Laternula: 2:35-40. L. truncata, Lyon-
sia hyalina: 3(1):104. Pecten: 1:13
(passim). Rhinoclava (Proclava):
2:1-20. Tridacna maxima: 1:18
(passim)

Faunal Replacement

Pelecypoda, Unspecified: 2:79

Fecundity

Cepaea nemoralis: 1:103

Feeding

Abra alba: 5(1):21-30 (passim).
Acanthodoris spp., Acteocina
canaliculata, Acteonia cocksi,
Adalaria: 5(2):197-214. A. proxima:
4(2):235; 5(2):197-214. Adipicola:
$1:23-34. Aegires spp., Aglaja
ocelligera, Aldaria modesta, Aldisa
binotata, A. cooperi, A. pikokai, A.
sanguinea, A. tara: 5(2):197-214.
Amygdalum, A. politum: $1:23-34.
Ancula pacifica, Anisodoris nobilis,
Antonietta luteorufa: 5(2):197-214.
Aplacophora: S1:23-34. Aplysia
juliana, Aplysiopsis smithi:
5(2):197-214. Archaeogastropoda:
$1:23-34. Archidoris odhneri, A.
pseudoargus, Armina californica, A.
maculata: 5(2):197-214. Ascobulla
ulla: 5(2):259-280. Astarte castanea:
5(1):21-30 (passim). Babaina:
5(2):197-214. Berthelinia caribbea:
5(2):197-214, 259-280. B. limax,
Berthella californica, Berthellina
citrina: 5(2):197-214. Bithynia ten-
taculata: 3(2):179-186. Bosellia
mimetica: 5(2):197-214, 259-280.
Bosellidae: 5(2):259-280. Brechites
penis: 5(1):21-30 (passim). Cadlina
laevis, C. modesta, Caliphylla medi-
terranea: 5(2):197-214. Caliphyllidae:
5(2):259-280. Calliopaea bellula,
Calma glaucoides, Calmella carolinii:
5(2):197-214. Calyptogena spp.:
$1:23-34. Calyptraea conica,

294 AMER.

Capulidae: S1:35-50. Casella ob-
soleta: 5(2):197-214. Cassis tuberosa:
$1:35-50. Catriona gymnota, C.
maua: 5(2):197-214. Chaetomorpha:
5(2):259-280. Chelidonura, Chromo-
doris spp.: 5(2):197-214. Codakia or-
bicularis: S1:23-34. Corbicula:
5(1):21-30 (passim). C. fluminea:
$2:167-178, 187-191, 219-222. Cor-
biculacea: 5(1):21-30 (passim).
Costasiella ocellifera: 5(2):197-214,
259-280. C. nonatoi, Costasiellidae:
5(2):259-280. Crassostrea virginica:
S3:41-49. Cratena peregrina:
5(2):197-214. Crenella: $1:23-34.
Crepidula fornicata: S1:35-50.
Crimora coneja, C. papillata,
Cumanotus beaumonti, Cuthona
spp.: 5(2):197-214. Cyerce antillensis:
5(2):259-280. C. cristallina:
5(2):197-214. Cymatium nicobaricum,
Cypraecassis testiculus: S1:35-50.
Dacrydium: $1:23-34. Dendrodoris
spp., Dendronotus spp., Dermato-
branchus Striatellus, Diaphana
californica, Dicata odhneri, Dirona
albolineata, D. aurantia, Discodoris
spp., Doridella obscura, D.
steinbergae, Doriopsilla pharpa,
Doris ocelligera, Doto spp., Elysia
spp.: 5(2):197-214, 259-280.
Embletonia pulchra faurei, Eolidina
mannarensis: 5(2):197-214. Epitonium
albidum: 1:1-12. Ercolania funerea, E.
fuscata: 5(2):197-214, 259-280.
Eubranchus spp., Facelina spp.:
5(2):197-214. Falcidens: S1:23-34.
Fimbria fimbriata: 5(1):21-30
(passim). Fioina pinnata, Flabella
spp., Flabellina affinis: 5(2):197-214.
Gastropoda, Unspecified: 4(1):114.
Glossodoris spp., Goniodoris
castanea, Gymnodoris striata, Hallaxa
chani, Haminoea spp., Hancockia
ucinata, Hermaea bifida, Hoplodoris
nodulosa: 5(2):197-214. Hydrobia
ulvae: S1:35-50. Hypselodoris ben-
netti, H. messinensis: 5(2):197-214.
Idasola, |. argentea, Lacuna coss-
manni: S1:23-34. Lalia cockerelli:
5(2):197-214. Lamellibranchia:
$1:23-34. Limapontia capitata:
5(2):197-214, 259-280. Limenanara
nodosa: 5(2):197-214. Lirularia, L.
lirulata: 4(1):109. Littorina littorea, L.
obtusata: 1:92. Lobiger serradifalci:
5(2):197-214. L. souverbiei:
5(2):259-280. Lucina spp., Lucinidae,
Lucinoma spp.: $1:23-34. Lunatia
lewisi: S1:35-50. Macoma balthica:
5(1):21-30 (passim). Melanochlamys
diomedea, Melibe fimbriata, M.
leonina: 5(2):197-214.

Mesogastropoda: S1:23-34. Miamira
sinuata: 5(2):197-214. Mitra idae:
1:91-92. Modiolus: S$1:23-34.
Mourgona germaineae: 5(2):259-280.
Musculium securis: 5(1):21-30
(passim). Musculus, Myrina,
Mytilidae, Neogastropoda,
Neomenia, Neomphalace, Neom-
phalidae, Neomphalus fretterae:
$1:23-34. Nucella lapillus: 2:63-73.
Okadaia elegans, Olea hansineensis,
Onchidoris spp.: 5(2):197-214. Oxynoe
antillarum: 5(2):259-280. O. azuro-
punctata: 5(2):197-214, 259-280.
Peltodoris atromaculata: 5(2):197-214.
Petromyzon marinus: 5(1):21-30
(passim). Phestilla melanobranchia,
P. sibogae, Phidiana crassicornis,
Philine gibba, Phyllaplysia engeli, P
taylori, Phylliroe bucephala, Piseino-
tecus sphaeriferus: 5(2):197-214.
Pisidium spp.: 5(1):21-30. Placida
cremoniana: 5(2):197-214. P den-
dritica, P kingstoni: 5(2):259-280. P
viridis, Platydoris scabra:
5(2):197-214. Pogonophora: $1:23-34.
Polycera quadrilineata, P zosterae,
Polycerella emertoni: 5(2):197-214.
Polymesoda (Geloina) erosa:
5(1):21-30 (passim). Precuthona
divae: 5(2):197-214. Prosobranchia,
Pseudomiltha: S1:23-34. Pteraeolidia
ianthina, Retusa obtusa, Rostanga
pulchra, Runcina ferruginea, R. seto-
ensis, Scyllaea pelagica, Sebradoris
crosslandi: 5(2):197-214. Simrothiella,
Simrothiellidae, Solemya (Acharax),
Solemya spp., Solemyidae: S1:23-34.
Sphaerium spp., S. corneum:
5(1):21-30 (passim). Stiliger fusco-
vittatus: 5(2):197-214. Stiligeridae:
5(2):259-280. Struthiolariidae:
$1:35-50. Tenellia pallida, Tergipes
tergipes, Tethys fimbria: 5(2):197-214.
Thais haemastoma: S$1:35-50. T.
haemastoma canaliculata: 2:63-73.
Thecacera pennifera, Thordisa filix,
Thorunna spp.: 5(2):197-214.
Thyrasira, Thyrasiridae: $1:23-34.
Trapania maculata, Tridachia
crispata, Triopha catalinae, Trippa
spongiosa, Tritonia diomeda, T. festiva:
5(2):197-214. IT hombergi: 4(2):235;
5(2):197-214. Tritoniopsis cincta:
5(2):197-214. Trochacea, Turridae:
$1:23-34. Turritellidae: S1:35-50. Um-
bonium : 4(1):109. Vesicomya spp.,
Vesicomyidae, Vestimentifera:
$1:23-34. Volvatella bermudae:
5(2):259-280

Feeding - Sediment Relationships

Pomacea paludosa, Stagnicola sp.:
1:97

MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Fertilization

Corbicula fluminea: 4(1):61-79

Filter Feeding

Calyptraea chinensis, Capulis
ungaris, Crepidula fornicata, Mytilus
edulis, Vivaparus viviparous:
3(2):179-186 (passim)

Filtration Rate

Dreissena polymorpha: S2:174
(passim)

Fisheries

Aequipecten circularis: 4(1):119.
Anadara spp.: 4(1):111. Busycon
spp.: 3(1):102. Cephalopoda, Un-
specified: 2:89. Chione cancellata:
4(1):111. Corbicula spp.: S2:1-5.
Crassostrea virginca: S3:1-4, 5-10,
11-16, 17-23. Haliotis roei: 3(1):97. Illex
spp., Loligo: S1:93-100. Mercenaria
mercenaria: 4(1):111; S3:41-49. Mya
arenaria: 4(1):120-121; S3:59-70. M.
truncata: 4(1):120-121. Nautilus
macromphalus: S1:93-100. Neotia
ponderosa: 4(1):111. Octopodidae:
$1:93-100. Octopus vulgaris: 4(2):240;
$1:93-100. Ostrea chilensis: S3:1-4.
O. edulis: S3:41-49. O. irridescens,
Pinctada mazatlanica: 4(1):119.
Polinices duplicatus: 4(1):111. Pro-
tothaca asperimma: 4(1):119.

Flight - Flash Coloration

Hexabranchus sanguineus, Ptero-
poda, Pycnopodia helianthoides, Tri-
tionia diomeda: 5(2):185-196

Food

Adalaria proxima: 5(2):197-214,
293-301. Aeolidia papillosa:
5(2):287-292. Alcyonium digitatum:
5(2):197-214. Aldaria modesta,
Alvania auberiana: 4(2):185-199. Ana-
baena: 4(1):81-88. A. oscillarioides:
$2:219-222. Ancylus fluviatilis:
3(2):243-265. Ankistrodesmus:
4(1):81-88, S2:219-222. Archidoris
monteryensis: 5(2):185-196. A.
pseudoargus: 5(2):185-196, 197-214.
Asterionella: S2:167-178. Aufwuchs:
3(2):169-177, 243-265. Bacillariophy-
caea: S2:167-178. Berthelinia carib-
bea: 5(2):197-214, 259-280. Berthelinia
limax, Bimeria: 5(2):197-214. Caecum
nitidum: 4(1):185-199. Catriona gym-
nota: 5(2):185-196. Caulerpa
okamurai: 5(2):197-214. C. verticilliata:
5(2):197-214, 259-280. Ceratium
hirundinella: S2:167-178. Chlamy-
domonas: 4(1):81-88. Chlorella:
4(1):81-88; S2:143-150, 167-178. C.
vulgaris: 3(2):179-186; S2:219-222.
Chlorophyceae, Chrysophyceae:
$2:167-178. Cliona celata:
5(2):185-196. Corbicula fluminea:
4(1):81-88; S2:143-150, 167-178,

AMER

219-222. Coryphella: 5(2):185-196.
Cuthona adyarensis: 5(2):197-214. C.
nana: 5(2):185-196, 287-292. Cyano-
phyceae, Dinophycea: S2:167-178.
Doridella obscura, D. steinbergae,
Electra crustulenta: 5(2):197-214. E.
pilosa: 4(1):103-104; 5(2):197-214,
293-301. Escherichia coli:
3(2):179-186. Eubranchus exiguus,
E. farrani: 5(2):197-214. Euglenophy-
caea: S2:167-178. Eurystomella
bilabriata, Facelina coronata:
5(2):185-196. Fragilaria: S2:167-178.
Granulina ovaliformis: 4(1):185-199.
Gymnodinium veneficum: S2:167-178.
Halichondria panicea: 5(2):185-196,
197-214. Halodule wrighti:
4(2):185-199. Hopkinsia rosacea:
5(2):185-196. Hydractinia echinata:
5(2):185-196, 287-292. Isochrysis:
$1:85-91. /. galbiana: 3(1):33-40;
4(1):81-88, 89-99. Jorunna tormentosa:
5(2):185-196. Kirchenpaueria pinnata,
Laomedea, L. loveni: 5(2):197-214.
Marginella aureocincta: 4(1):185-199.
Melosira: S2:167-178. Membranipora
villosa: 5(2):197-214. Monochrysis
lutheri: 3(1):33-40; 4(1):89-99. Nitz-
schia actinastroides: 3(2):151-168.
Onchidoris muricata: 5(2):197-214,
293-301. Oplitaspongia pennata:
5(2):197-214. Phestilla melano-
branchia: 5(2):185-196, 197-214. P
sibogae: 5(2):197-214. Porites
somaliensis: 5(2):197-214. Puperita
pupa: 4(2):185-199. Rissoella
caribaea, Rissoina bryerea, R. cates-
byana: 4(2):185-199. Rostanga
pulchra: 5(2):197-214. Scenedesmus:
4(1):81-88; S2:143-150. Skeletonema
costatum: 4(1):81-88. Smaragdia
viridis viridemaris: 4(2):185-199.
Stephanodiscus, Synedra:
$2:167-178. Tenellia pallida:
5(2):197-214. Thalassia testudinum:
4(2):185-199. Tricolia spp.:
4(2):185-199. Tritonia diomeda, T.
hombergi, Tubastraea coccinea:
5(2):197-214. Turbinaria: 5(2):185-196.
Vaucheria, Virgularia: 5(2):197-214.
Zebina browniana: 4(2):185-199

Food, human use as
Unionidae: 1:31-34

Foot
Busycon contrarum: 4(1):110. Cor-
bicula fluminea: 2:87. Gastropoda,
Unspecified: 4(2):243

Founder Effect
Cepaea sp.: 1:103

Fortuitous Coloration
Opisthobranchia: 5(2):185-196

G-Band, chromosome

Biomphalaria glabrata, B. straminea:

1:106-107

Gametogenesis
Anodonta imbecilis: 4(1):117;
4(2):231. Corbicula fluminea:
4(1):61-79. Elliptio icterina, Villosa
villosa: 4(1)117; 4(2):231

Garstang Torsion Theory
Gastropoda, Unspecified: 1:89

Gene Flow
Partula taeniata: 1:103-104

Genetics
Amblemini: 1:109-110. Ancylus fluvi-
atilis: 5(1):105-124. Arianta ar-
bustorum: 1:103. Arion spp.: 1:24,
110. Ashmunella spp.: 1:21-26, 106.
Biomphalaria spp.: 1:106, 1:106-107,
1:107. Bradybaenidae: 2:97. Bulinus
spp.: 1:106-107. Cepaea spp.: 1:103,
1:107-108. Corbicula: 1:96; S2:83-88,
124-132. C. fluminea: S2:89-94.
Crassostrea, C. rhizophorae, C.
virginica: 1:108-109. Crepidula spp.:
1:110. Deroceras laeve: 1:23
(passim), 1:110. D. reticulatum: 1:110.
Elliptio spp., Elliptoideus, Fusconaia:
1:109-110. Goniobasis proxima: 1:105;
3(1):99-100. Helix aspersa: 1:24
(passim). Lampsilis: 1:109-110. Liguus
spp.: 5(2):153-157. Littorina: 1:108-109.
Lymnaea (Stagnicola) elodes:
5(1):105-124; 6(1):9-17. Macoma:
1:109-110. M. balthica: 1:90.
Megalonaias: 1:109-110. Megapallifera
mutabilis, Meghimatium: 4(2):238.
Mercenaria mercenaria: 1:107.
Mesoaon, M. zaletus: 2:97-98.
Modiolus, Mytilus: 1:108-109. M.
desolationis: 1:105-106. M. edulis, M.
galloprovincialis: 1:105-106, 1:108.
Nucella emarginata: 1:105. Ostrea
edulis: 1:105-106. Pallifera: 4(2):238.
Partula spp.: 1:103-104. Phylomyci-
dae, Phylomycus carolinianus, P.
togatus: 4(2):238. Pisidium caser-
tanum: 5(1):49-64. Quadrula, Quin-
cuncina: 1:109-110. Rumina decollata:
1:23 (passim). Sphaerium Sstriatinum:
5(1):49-64 (passim). Thais
emarginata: 1:105. Theba pisana:
1:104, 104-105. Triodopsis: 2:97-98

Gills
Chaetopleura apiculata: 6(1):69-78

Gizzard Stones
Pomacea paludosa, Stagnicola sp.:
1:97

Glands, Digestive
Corbicula fluminea: 3(1):101;
4(1):115-116

Glands, Gill
Archidoris pseudoargus, Peltodoris
atromaculata: 4(2):232

Glands, Hypobranchial
Nucella lapillus: S1:35-50

. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 295

Glands, Mantle
Clavagella australis, Clavagellidae,
Cleidothaeridae, Cuspidariidae, En-
todesma: S1:35-50. Laternulidae:
2:35-40. Lyonsiidae, Myochamidae,
Mytilimera nutalli, Pandoridae,
Parilimya fragilis, Parilimyidae, Peri-
ploma fragile, P (Offadesma) angasi,
Periplomatidae, Pholadomya can-
dida, Pholadomyidae, Thracia
phaseolina, Thraciidae, Verticor-
diidae: S1:35-50

Glochidia
Alasmidonta marginata: A. viridis,
Amblema plicata plicata,
Amblemidae, Anodonta sp., Anodon-
toides: 4(1):117-118. Elliptio:
5(2):125-128. Epioblasma, Fusconaia
ebena, Lampsilis: 4(1):117-118. L. hig-
ginsi: 6(1):39-43. Lasmigona spp.,
Leptodea, Magnonaias nervosa:
4(1):117-118. Margaritifera laevis, M.
margaritifera: 5(2):125-128. Obovaria,
Pegias, Potamilus, Ptychobranchus,
Quaadtrula cylindrica cylindrica, Q.
pustulosa pustulosa, Strophitus un-
dulatus tennessensis, Tritogonia,
Villosa: 4(1):117-118

Glochidial Host
Onchorhyncus kisutch, O. tshawyt-
scha, Salmo salar (all for Margaritifera
margaritifera): 5(2):125-128 (passim).
S. trutta (for Margaritifera margari-
tifera): 5(2):125-128

Growth
Ancylus fluviatilis: 5(1):105-124.
Australorbis glabratus: 3(2):213-221.
Cepaea nemoralis: 1:103. Cistopus
indicus: 6(2):207-211. Corbicula:
$2:41-45, 47-52, 53-58. C. fluminea:
3(1):100; 4(1):81-88; S2:69-81, 133-142,
143-150, 151-166, 167-178, 211-218,
231-239. Crassostrea virginica:
$3:41-49. Elliptio icterina: 1:95.
Epitonium albidum: 1:1-12. Ferrissia
rivularis: 5(1):105-124 (passim). Gastro-
poda, Unspecified: 2:80-81. Hapaloch-
laena maculosa: 6(2):207-211.
Laevapex fuscus 5(1):105-124
(passim). Littorina littorea: 1:92;
5(1):105-124. L. obtusata: 1:92. Loligo
opalescens: 2:93. Lymnaea
(Stagnicola) elodes: 3(2):143-150,
213-221; 5(1):105-124. L. palustris:
3(2):213-221. Macoma balthica: 1:90;
3(2):213-221. Margaritifera margariti-
fera: 5(1):105-124 (passim).
Mercenaria mercenaria: 4(2):149-155.
Musculium partumeium: 5(1):49-64
(passim). M. securis: 5(1):49-64
(passim). Mya arenaria, M. truncata:
4(1):120-121. Nucella lapulus: 4(1):110.
Octopus spp.: 2:92; 6(2):207-211.

296 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Pisidium casertanum: 5(1):49-64.
Placopecten magellanicus: 6(1):1-8.
Planorbis corneus: 3(2):213-221.
Pteroctopus tetracirrhus, Robsonella
fontanianus, Scaeurgus patagiatus,
S. unicirrhus: 6(2):207-211.
Sinonovacula: 5(2):159-164. Villosa
villosa: 1:95. Viviparus georgianus:
3(2):268

Growth Bands, External
Lasmigona subviridis, Medionidus
conradicus, Pleurobema oviforme,
Villosa vanuxemi: 6(2):179-188

Gugler, Carl W.
Obituary: 3(1):83-84

Habitat
Batillaria minimum: 2:1-20.
Buchanania, B. onchidioides:
2:21-34. Cerithidea spp.: 2:1-20.
Epitonium albidum: 1:1-12. Fissurelli-
dea spp., Pupillaea spp.: 2:21-34

Habitat Distribution
Unionidae: 1:61-68

Habitat Stability
Cepaea sp.: 1:103

Hatching Size
Helisoma trivolvis: 4(2):229

Heterochrony
Corbicula fluminea: 4(1):61-79

Histochemistry
Aeolidia papillosa: 4(2):205-216.
Boonea impressa: 3(1):97. Cionella
lubrica: 3(1):27-32. Clavagella
australis: S1:35-50. Coryphella
salmonacea, Hermissenda crassi-
cornis: 4(2):205-216. Odostomia im-
pressa: 3(1):97

Hormones
Cryptozona belangeri: 4(1):115;
4(2):237

Hybridization
Crassostrea rhizophorae, C. virginica:
1:108

Hydrothermal Vents
Adipicola, Amygdalum, A. politum,
Aplacophora, Archaeogastropoda,
Calyptogena: S1:23-34. C. magnifica:
1:101; 4(1):49-54; S1:23-34. C. ponder-
osa, Codakia orbicularis, Crenella,
Dacrydium, Falcidens, Idasola, |. ar-
gentea, Lacuna cossmanni, Lamelli-
branchia, Lucina atlantis, L. (Linga)
pennsylvanica, L. (Phacoides) pectina-
tus, Lucinidae, Lucinoma, L. atlantis,
L. filosa, Mesogastropoda, Modiolus,
Musculus, Myrina: S1:23-24.
Mytilidae: 1:101; 3(1):95; S1:23-34.
Neogastropoda, Neomenia, Neom-
phalace, Neomphalidae, Neom-
phalus fretterae, Patellidae,
Pogonophora, Prosobranchia,
Pseudomiltha, Simrothiella, Simrothi-
ellidae, Solemya (Acharax) caribbaea,

S. (Acharax) johnsoni, S. agassizi, S.
velum, Solemyidae, Thyrasira,
Thyrasiridae: S1:23-34. Trochacea:
3(1):104; S1:23-34. Trochidae: 3(1):95.
Turridae, Vesicomya, V. caudata, V.
cordata: $1:23-34. Vesicomyidae:
1:101; 3(1):95-96; S1:23-34.
Vestimentifera: S1:23-34

Immunology
Amoeba proteus, Biomphalaria
glabrata, Chilomonas, Colpidium,
Crassostrea virginica, Daphnia, Liolo-
phura gaimardi, Monas, Mya
arenaria, Mytilus edulis, Periplaneta
americana, Schistosoma mansoni:
$1:79-83

Inbreeding
Littorina, Macoma, Mytilus: 1:108-109

Infection
Bankia gouldi: S1:101-109. Crasso-
strea virginica: S1:101-109 (passim);
$3:5-10, 17-23. Boveria terediniai:
$1:101-109. B. zeukevitchi: S1:101-109.
Haplosporidia nelsoni: S3:5-10. Hap-
losporidium: $1:101-109; S3:5-10. H.
costalis: S3:59-70. H. (Minchinia)
nelsoni: S3:17-23, 59-70. Octopus
briareus, O. joubini: 2:93-94. Teredo
spp.: S1:101-109. Vibrio spp.: 2:93-94

Invasion History
Corbicula fluminea: 1:100; S2:1-5,
7-39

Iridophores
Lolliguncula brevis: 2:91

Isolation, Genetic
Partula mooreana, P. suturalis, P
taeniata: 1:103-104

Karyotype
Ashmunella lenticula, A. proxima
albicaudata: 1:106. Bellamya spp.:
4(1):107. Biomphalaria glabrata, B.
straminea: 1:106-107. Bradybaena
similaris, B. (Acusta) despecta siebol-
diana: 2:97. Caelatura: 4(1):107.
Crassostrea virginica: 1:105-106.
Euhadra: 2:97. Megapallifera
mutabilis: 4(2):238. Mytilus spp.:
1:105-106. Neothauma tanganyicense:
4(1):107. Ostrea edulis: 1:105-106.
Phylomycidae, Phylomycus carolini-
anus, P. togatus: 4(2):238. Unionidae,
Unspecified: 2:86-87. Viviparidae:
3(1):107

Kidney
Aciculidae, Ampullariidae, Assiminei-
dae, Bithyniidae, Buccinum undatum,
Cerithiidae, Cyclophoridae, Deroceras
reticulatum, Haliotis corrugata, H.
rufescens, Helicinidae, Helix
pomatia, Hydrobiidae, Hydrocenidae,
Limax pseudoflavus, Littorina irrorata,
Lymnaea stagnalis, Marisa cornuari-
etis, Melaniidae, Melanoposidae,

Mesogastropoda, Nerita fulgurans,
Neritacea, Neritidae, Neritina
latissima, Patella vulgata, Pleuroceri-
dae, Pomacea lineata, Potamopyrgus
jenkinsii, Rissoacea, Rissoidae,
Strombus gigas, Syrnolopsidae,
Thiaridae: 3(2):223-231. Tridacna sp.:
2:83. Valvatacea, Valvatidae,
Viviparacea, Viviparidae, Viviparus
spp.: 3(2):223-231

Kidney Function
Tridacna sp.: 2:83

Laboratory Culture
Biomphalaria glabrata: 3(1):89-90. II-
lex spp., Loligo, Nautilus macrom-
phalus, Octopodidae: $1:93-100. Oc-
topus dofleini martini: 4(2):241. O.
vulgaris: S1:93-100

Larval Settlement
Bankia gouldi: 4(1):89-99. Chrysaora
quinquecirrha: S3:59-70. Crassostrea
virginica: 4(1):101; S3:59-70. Diadumene
leucolena, Mnemiopsis leidyi: S3:59-70.
Teredo bartschi, T. navalis: 4(1):89-99

Larvae
Acanthodoris spp.: 5(2):197-214.
Aclididae, Aclis, Acochlidiacea,
Acteocina sp.: $1:1-22. A.
canaliculata: 5(2):197-214.
Acteocinidae, Acteon: $1:1-22. Ac-
teonia cocksi, Adalaria: 5(2):197-214.
A. proxima: 4(1):103-104;
5(2):197-214, 293-301. Aegires spp.:
5(2):197-214. Aglaja: $1:1-22. A.
ocelligera: 5(2):197-214. Aglajidae,
Akera, Akeridae: $1:1-22. Aldaria
modesta, Aldisa spp.: 5(2):197-214.
Allogastropda, Amaea: $1:1-22. Am-
nicola winkleyi: 4(1):101-102. Amphi-
bola, Amphibolidae, Anaspidea:
$1:1-22. Ancula pacifica:
5(2):197-214. Angutispira: S1:1-22.
Anisodoris nobilis, Antonietta
luteorufa: 5(2):197-214. Aplysia sp.:
$1:1-22. A. juliana: 5(2):197-214.
Aplysiidae, Aplysiomorpha: S$1:1-22.
Aplysiopsis smithi: 5(2):197-214. Arca
noae: S1:59-78. Archidoris odhneri:
5(2):197-214. A. pseudoargus:
4(1):103-104; 5(2):197-214. Architec-
tonicacea, Architectonicidae: S1:1-22.
Arctica islandica: $1:59-78; S3:51-57.
Argopecten irradians: $1:59-78. Ar-
mina californica, A. maculata:
5(2):197-214. Ascoglossa: S$1:1-22.
Astarte castanea: S1:59-78.
Atyidae: S1:1-22. Babaina: 5(2):197-214.
Bankia gouldi: 4(1):89-99.
Basommatophora, Berthelinia:
S$1:1-22. B. caribbea, B. limax:
5(2):197-214. Berthella: S1:1-22. B.
californica: 5(2):197-214. Berthellina:
$1:1-22. B. citrina: 5(2):197-214.

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 297

Bittum alternatum: $1:85-91.
Bivalvia, Unspecified: 4(1):102-103.
Blauneria, Boonea: S1:1-22. Bosellia
mimetica: 5(2):197-214. Bulla,
Bullidae, Bullina, Bullomorpha:
$1:1-22. Cadlina laevis: 4(1):103-104;
5(2):197-214. C. modesta, Caliphylla
mediterranea, Calliopaea bellula,
Calma glaucoides, Calmella carolinii:
5(2):197-214. Calyptogena magnifica:
4(1):49-54. Calyptraeidae: $1:1-22.
Campanile, Campanilidae, Cary-
chium: $1:1-22. Casella obsoleta,
Catriona gymnota, C. maua:
5(2):197-214. Cephalaspidae, Cerithi-
opsacea, Cerithiopsidae: S1:1-22.
Chelidonura: 5(2):197-214; $1:1-22.
Chilina, Chilinidae: S1:1-22. Chromo-
doris spp.: 5(2):197-214. Chrysallida:
$1:1-22. Cincinnatia winkleyi:
4(1):101-102. Corbicula: S2:41-45,
47-52, 53-58, 63-67, 83-88, 95-98. C.
fluminea: 2:87; 4(1):61-79, 81-88;
S2:69-81, 99-111, 151-166, 193-201.
Costasiella ocellifera: 5(2):197-214.
Couthouyella: S1:1-22. Crassostrea
virginica: 4(1):101; S1:59-78; S3:1-4,
5-10, 11-16, 25-29, 31-36, 41-49, 59-70,
71-75. Cratena peregrina:
5(2):197-214. Crepidula spp.:
$1:85-91. Crimora coneja, C. papil-
lata, Cumanotus beaumonti, Cuthona
spp.: 5(2):197-214. Cyclocardia
borealis: S1:59-78. Cyclophoridae,
Cyclostremella, Cyclostremelidae:
$1:1-22. Cyerce cristallina:
5(2):197-214. Cylinchna: $1:1-22.
Cylinchnella canaliculata: 1:91. Cylin-
drobulla, Cylindrobullidae: S1:1-22.
Cymatium parthenopeum: S1:85-91.
Cymbulia, Cymbuliidae: $1:1-22.
Dendrodoris spp., Dendronotus spp.,
Dermatobranchus Striatellus:
5(2):197-214. Detracia, Diaphana:
$1:1-22. D. californica: 5(2):197-214.
Diaphanidae: $1:1-22. Dicta odhneri,
Dirona albolineata, D. aurantia,
Discodoris spp.: 5(2):197-214.
Docoglossa: $1:1-22. Doridella
obscura, D. steinbergae, Doriopsilla
pharpa, Doris ocelligera, Doto spp.:
5(2):197-214. Ebala, Ellobiidae,
Ellobium, Elysia: S1: 1-22. E. spp.:
5(2):197-214. Elysiidae: $1:1-22.
Embletonia pulchra faurei:
5(2):197-214. Ensis directus: S1:59-78.
Entomotaeniata: S1:1-22. Eolidina
mannarensis: 5(2):197-214. Epitoni-
acea, Epitoniidae, Epitonium, E.
albidum: 1:1-12. Ercolania funerea, E.

fuscata, Eubranchus spp.: 5(2):197-214.

Eulimacea, Eulimidae, Euthyneura:
$1:1-22. Facelina spp.: 5(2):197-214.

Fargoa bartschi: $1:1-22. Fiona pin-
nata, Flabella spp., Flabellina affinis:
5(2):197-214. Gastropoda, Unspeci-
fied: 4(1):102-103, 103. Gegania:
$1:1-22. Geukensia demissa:
$1:59-78. Gleba: S1:1-22. Glossodoris
spp., Goniodoris castanea, Gym-
nodoris striata: 5(2):197-214. Gym-
nosomata: $1:1-22. Hallaxa chani:
5(2):197-214. Haminoea: $1:1-22. H.
spp., Hancockia ucinata:
5(2):197-214. Hedylopsidae, Hedylop-
sis, Heliaucus, H. cylindricus, H. per-
reieri: S1:1-22. Hermaea bifida:
5(2):197-214. Heterobranchia, Hetero-
gastropoda, Heteroglossa: $1:1-22.
Hoplodoris nodulosa: 5(2):197-214.
Hydatina, Hydatinidae: S1:1-22.
Hyarobia truncata: 4(1):101-102.
Hypselodoris bennetti, H. messinen-
SiS: 5(2):197-214. Illex illecebrosus:
4(2):240-241. Janthina spp., Jan-
thinidae: S1:1-22. Jorunna tormen-
tosa: 4(1):103-104. Juliidae: $1:1-22.
Lalia cockerelli: 5(2):197-214. Latia,
Latiidae, Leucophytia, Limacinidae,
Limapontia: $1:1-22. Limapontia
capitata: 5(2):197-214. Limapontiidae:
$1:1-22. Limenandra nodosa:
5(2):197-214. Littorina: S1:1-22.
Lobiger serradifalci: 5(2):197-214.
Lymacina: $1:1-22. Macoma balthica:
$1:59-78. Mathilda, Mathildidae,
Maxacteon, Melampidae, Melampus:
$1:1-22. Melanochlamys diomedea,
Melibe fimbriata, M. leonina:
5(2):197-214. Mellanella spp.: 2:83.
Mercenaria mercenaria: S1:59-78.
Mesogastropoda: S$1:1-22. Miamira
sinuata: 5(2):197-214. Micromelo:
$1:1-22. Modiolus modiolus: 4(1):104;
$1:59-78. Mopalia mucosa: S$1:85-91.
Mulnia spp.: 4(1):104. M. lateralis:
$1:59-78. Muricidae: S1:1-22. Mya
arenaria, Mytilus californianus:
$1:59-78. M. edulis: 4(1):104;
$1:59-78, 85-91. Myxa, Neogastro-
poda, Notaspidae, Nudibranchia,
Odostomia: $1:1-22. Okadaia
elegans, Olea hansineensis:
5(2):197-214. Omalogyra, Onchidella,
Onchidiidae, Onchidium: $1:1-22.
Onchidoris spp.: 4(1):103-104;
5(2):197-214, 293-301. Opistho-
branchia: S1:1-22. Ostrea chilensis:
$3:1-4. Otina, Otinidae, Ovatella, Ox-
ynidae, Oxynoe: $1:1-22. O. azuro-
punctata, Peltodoris atromaculata:
5(2):197-214. Peracle, Peraclidae,
Phanerophthalmus: S1:1-22. Phestilla
melanobranchia, P. sibogae, Phidi-
ana crassicornis: 5(2):197-214. Philine:
$1:1-22. P gibba: 5(2):197-214.

Philinidae, Philinoglossa, Philino-
glossidae, Philippia, Pholadidae:
$1:59-78. Phyllaplysia engeli, P. taylori,
Phylliroe bucephala, Piseinotecus
sphaeriferus, Placida cremoniana, P
viridis: 5(2):197-214. Placopecten
magellanicus: 4(1):104; S1:59-78.
Planorbidae: $1:1-22. Platydoris
scabra: 5(2):197-214. Pleurobranchi-
dae, Pleurobranchomorpha, Pleuro-
branchus: $1:1-22. Polycera quaarili-
neata, P zosterae, Polycerella emer-
toni, Precuthona divae: 5(2):197-214.
Prosobranchia, Pseudomalaxis,
Pseudoskenella, Ptenoglossa: $1:1-22.
Pteraeolidia ianthina: 5(2):197-214.
Pulmonata, Pupa, Purpura patula,
Pyramidella crenulata, Pyramidella-
cea, Pyramidellidae, Pythia, Radix:
$1:1-22. Retusa obtusa: 5(2):197-214.
Retusidae, Retussa, Ringicula, Ringi--
culidae, Rissoella, Rissoellidae: S1:1-22.
Rostanga pulchra: 5(2):197-214.
Roxania: S1:1-22. Runcina fer-
ruginea, R. setoensis: 5(2):197-214.
Sacoglossa, Salinator, Sayella,
Scaphander, Scaphanderidae: $1:1-22.
Scyllaea pelagica, Sebradoris cross-
landi: 5(2):197-214. Siphonaria,
Siphonariidae, Smaragdinella: $1:1-22.
Spisula solidissima: $1:59-78. Spur-
winkia salsa: 4(1):101-102. Stiligar:
$1:1-22. Stiliger fuscovittatus:
5(2):197-214. Stiligeridae, Systellom-
matophor: S$1:1-22. Tenellia pallida:
5(2):197-214. Teredo bartschi, T.
navalis: 4(1):89-99. Tergipes tergipes,
Tethys fimbria: 5(2):197-214. Thais
haemastoma canaliculata: 6(2):189-197.
Thecacera pennifera: 5(2):197-214.
Thecosomata: S1:1-22. Thordisa filix,
Thorunna spp.: 5(2):197-214. Toledella:
$1:1-22. Transennella tantilla: 2:94.
Trapania maculata, Tridachia cris-
pata, Triopha catalinae: 5(2):197-214.
Triophridae: S1:1-22. Trippa
spongiosa, Tritonia diomeda, T. festiva:
5(2):197-214. T. hombergi:
4(1):103-104; 5(2):197-214. Tritoniopsis
cincta: 5(2):197-214. Trochidae, Tur-
bonilla, T. vineae, Turritellidae, Um-
braculidae, Umbraculum: $1:1-22.
Unionidae, Unspecified: 4(1):101. Val-
vata, Valvatacea, Valvatidae, Veron-
icellidae, Volvatella, Volvatellidae,
Williamia: $1:1-22

Learning

Limax maxima: 2:78

Life Cycle

Corbicula fluminea: 1:96. Littorina
saxatilis: 1:92-93. Mazatlania aciculata:
1:92. Triodopsis tridentata triden-
tata: 1:98

298 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Ligament
Bivalvia, general: 4(1):111-112

Light Emitting Diodes: 1:89

Locomotion
Aplacophora: S$1:35-50. Aplysia cali-
fornica: 2:78. Ariolimax colmbianus,
Cionella lubrica: $1:35-50. Corbicula
fluminea: 2:87; S2:187-191. Gastro-
poda, Unspecified: 4(2):243. Helix
aspersa: $1:35-50. Illex illecebrosus:
4(1):55-60. Lyonsia: S1:35-50.
Nautilus: 4(2):239-240. Patella
vulgata: S1:35-50. Periploma:
2:35-40. Polyplacophora, Unela
nahantensis: S1:35-50

Mantle
Corbiculacea, Corbiculidae: 4(1):116.
Crassostrea rhizophorae: 1:102.
Laternulidae: 2:35-40. Mytilus:
5(2):159-164 (passim). Pandoracea:
$1:35-50. Perna viridis: 5(2):159-164.
Pisidiidae: 4(1):116

Marginal Denticles, Homology
Hyotissa: 1:90

Mechanoreceptors
Navanax inermis: 1:13 (passim)

Metabolism
Musculium spp.: 3(2):187-200.
Physella virgata virgata: 3(2):243-265.
Pisidium spp., Sphaerium spp.:
3(2):187-200

Microstructure
Acanthochiton fascicularis: 6(1):141-151.
Aeolidia papillosa: 4(2):205-216.
Anguispira alternata: 4(2):237.
Chaetopleura lurida, C. peruviana:
6(1):141-151. Chiton olivaceus:
6(1):131-139, 141-151. Coryphella
salmonacea: 4(2):205-216. Eudoxo-
chiton nobilis: 6(1):141-151. Her-
missenda crassicornis: 4(2):205-216.
Ischnochiton herdmani, Katharina
tunicata: 6(1):141-151. Lasmigona
costata: 2:35-40. Lepidochitona
cinerea, L. dentiens, Lepidozona
retiporosus: 6(1):141-151. Lepido-
pleurus cajetanus: 6(1):141-151,
153-159. Mopalia spp., Placiphorella
velata, Plaxiphora obtecta, Tonicella
insignis: 6(1):141-151

Microstructure, Periostracum
See Periostracum Microstructure

Microstructure, Shell
See Shell Microstructure

Migration
Eledone cirrhosa: 6(1):45-48 (passim)

Mimicry
Aegires sublaevis, Aeolidia papillosa,
Aeolidiella glauca, A. sanguinea,
Aeolidiopsis, Aldisa, A. banyulensis,
Anisodoris, Aplysia spp.: A. parvula,
Archidoris, A. monteryensis, A.
pseudoargus, Ataagena, Bursatella:

5(2):185-196. Catriona gymnota:
5(2):185-196, 287-292. Cimora coneja,
Collembola, Coryphella: 5(2):185-196.
C. spp., Cuthona spp.: 5(2):185-196,
287-292. Cuthona poritophages, Den-
drodoris, Discodoris, Dondice
paguerensis, Dolabrifera, Doridella
obscura, D. steinbergae, Dorido-
morpha gardineri, Doriopsilla, D.
pharpa, Doris, Elysia arena:
5(2):185-196. Eubranchus:
5(2):243-258. E. exiguus: 5(2):185-196.
E. sanjuanensis, E. tricolor, Facelina
bostoniensis: 5(2):287-292. F. cor-
onata, Favorinus branchialis,
Gasterosteus aculeatus, Glaucus
atlanticus, Haminoea navicula,
Haplochromis burtoni, Hopkinsia
rosacea, Jorunna tormentosa, Laicus
argentatus: 5(2):185-196. Lalia
cockerelli: 5(2):287-292. Laomedea,
Obelia, Phestilla spp., Phyllaplysia
zostericola, Phyllodesmium spp.,
Pinufius rebus: 5(2):185-196.
Rostanga, R. pulchra, R. rubra:
5(2):185-196. Setoaeolis pilata:
5(2):287-292. Spurilla neapolitana,
Tergipes tergipes: 5(2):185-196.
Triopha catalinae: 5(2):287-292.
Tritonia nilsodhneri: 5(2):243-258

Mineralization, Periostracum
See Periostracum Mineralization

Mineralization, Shell
See Shell Mineralization

Modelling
Gastropod Growth: 2:80-81

Morph Frequencies
Arianta arbustorum: 1:103

Morphogenesis
Cephalopoda: 6(2):207-211

Morphology
Amplirhagada: 1:98-99. Argopecten

irradians: S1:59-78. Boonea impressa:

3(1):97. Colisella pelta: 2:80.
Curvemysella, C. paula: 1:90-91.
Epitonium albidum: 1:1-12. Gastro-
poda, Unspecified: 4(1):114. Haliotis
cracherodii: 4(2):233-234. Helisoma:
$1:51-58. Illex illecebrosus: 2:51-56.
Lampsilis altilis: 1:94. L. higginsi:
6(1):39-43. L. perovalis: 1:94. Ligumia
subrostrata: $1:51-58. Liguus spp.:
§(2):153-157. Littorina obtusata:
4(1):108. Lymnaea stagnalis:
$1:51-58. Micrarionta opuntia, M.
sodalis: 3(1):98. Mytilus edulis, M.
galloprovincialis: 1:108. Nautilus:
$1:51-58. Odostomia impressa:
3(1):97. Perna viridis: 5(2):159-164.
Pomacea paludosa: S1:51-58. Sep-
tifer: 5(2):159-164. Symplectoteuthis
oualaniensis: 2:51-56. Westraltrachia:
1:98-99

Morphology, Functional
Anomia simplex: 1:101-102; 2:41-50.
Corbicula fluminea: 1:13-20.
Lithophaga nigra: 1:101

Morphology, Shell
See Shell Morphology

Morphometrics
Ancylus fluviatilis: 5(1):105-124.
Campeloma geniculum, C. parthenum:
3(1):99. Cisopus indicus: 6(2):207-211.
Elliptio angustata, E. lanceolata:
1:95. Fontelicella: 4(2):243. Hapa-
lochlaena maculosa: 6(2):207-211.
Hydrobiidae, Lepidochitona dentiens:
4(2):243. Loligo sanpaulensis:
6(2):213-217. Lymnaea (Stagnicola)
elodes: 5(1):105-124. Octopus spp.,
Pteroctopus tetracirrhus, Robsonella
fontanianus, Scaeurgus patagiatus,
S. unicirrhus: 6(2):207-211.

Mortality
Arianta arbustorum, Cepaea hortensis:
1:103. Corbicula: S2:89-94. C.
fluminea: 3(1):94; S2:89-94. Crasso-
strea virginica: S3:5-10. Mercenaria
mercenaria: 4(2):149-155. Octopus
briareus, O. joubini: 2:93-94

Mucins
Mollusca, general: S1:35-50

Miullerian Mimicry
Opisthobranchia: 5(2):185-196

Multivariate Analysis
Goniobasis proxima: 1:105

Muscle
Anguispira alternata: 3(1):27-32
(passim). Anodonta spp.: 2:82. Arion
ater, Busycon canaliculatum, B. carica:
3(1):27-32 (passim). Cionella lubrica:
3(1):27-32. Lasmigona costata: 2:35-40.
Leucophyta bidentata: 3(1):27-32.
Lymnaea peregra, Melampus bidenta-
tus, Ofina otis, Pisania maculosa,
Pomatias elegans, Radis peregia, Unela
nahanensis: 3(1):27-32 (passim)

Museum
Canadian National Mollusc Collec-
tion: 2:81

Nephrolith
Tridacna spp.: 2:83

Nerves
Corbicula fluminea: 1:13-20

Nervous System
Batillaria minima, Cerithidea
scalariformis: 2:1-20

Neuropeptides
Aplysia spp.: 2:78

Neurophysiology
Aplysia spp., Limax maxima, Tritonia
diomeda: 2:78

Nucleic Acids
Cionella lubrica: 3(1):27-32

Odontophore Cartilage
Thais haemostoma canaliculata: 2:63-73

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 Paeye)

Old, William Erwood, Jr.
New Molluscan Taxa: 1:76. Obituary:
1:75-78. Publications: 1:76-77.
Species named in honor of: 1:76

Operculum
Cerithidea scalariformis: 2:1-20

Oral Shield
Chaetoderma, Falcidens, Limifossor,
Metachaetoderma, Prochaetoderma,
Scutopus, S. megaradulatus: 6(1):57-68

Organ Growth Rates
Elliptio lanceolata: 1:94-95

Osphradium
Camanillidae, Cerithidea, Diastoma-
tidae: 2:1-20. Lampsilis ventricosa:
1:18 (passim). Lymnaea stagnalis:
1:13 (passim). Modulidae, Planaxis:
2:1-20

Oxygen Tension
Gastropoda, Unspecified: 2:87-88

Paleobiogeography
Pelecypoda, Unspecified: 2:79

Paleontology
Acteocina, A. canaliculata, Acteon
wetherilli: 4(1):39-42, Amianthus:
4(1):1-12. Ammonites: 2:79. Anadara
(Cunearca) nux, A. (Esmerarca):
4(1):1-12. Ancistrobasis: 1:92.
Balanus spp.: 4(1):39-42. Bellamya
spp.: 4(1):107. Calliostoma hannibali,
Calyptraea: 4(1):1-12. Camaenidae:
3(1):8 (passim). Cancellaria (Pyruclia)
diadela: 2:84-85. Caracolus: 3(1):8
(passim). Cardita (Cardites): 4(1):1-12.
Cerithidea spp.: 2:1-20. Cerithium,
Chione spp., Choromytilus pallio-
punctatus: 4(1):1-12. Columbellidae:
3(1):96. Concavus, C. finchii, Conus
marylandicus: 4(1):39-42. Crassatella
ponderosa, C. vadosa, Crassatellidae:
4(2):238. Crassilabrum wittichi,
Crassispira starri, Crepidula: 4(1):1-12.
C. costata: 4(1):39-42. Crucibulum in-
erme, C. scutellatum, Cyclinella:
4(1):1-12. Cymia chelonia: 2:84-85.
Cymia heimi, Cypraea amandusi,
Divalinga comis, Drillia (Clathrodrillia):
4(1):1-12. Fusinus pumilus:
4(1):39-42. Fusus kingii: 4(2):236.
Gastropoda, Unspecified: 2:80-81.
Helminthoglyptidae, Hemitrochus:
1:97 (passim). Heteroterma, Hindsia
nodulosa: 4(2):236. Hipponix
pilosus: 4(1):1-12. Juliamitrella: 3(1):96.
Knefastia, Lucina (Lucinisca): 4(1):1-12.
Lysinoe, L. ghiesbreghti: 3(1):102-103.
Macron hartmanni: 4(1):1-12. Mactra
spp.: 4(1):39-42. Melongena melon-
gena: 4(1):1-12. M. melongena con-
sors: 2:84-85; 4(1):1-12. Mercenaria:
3(1):85-88. Micrarionta opuntia, M.
sodalis: 3(1):98; 4(2):237. Miliola mary-
landica, Mitrella communis: 4(1):39-42.

Mollusca, Unspecified: 2:79, 84;
3(1):96-97; 4(1):115; 4(2):238-239.
Mulinia lateralis: 4(1):39-42. Mytilus
canoasensis vidali, Nassarius versi-
color: 4(1):1-12. Nekewis: 4(2):236.
Nerita funiculata, Neverita (Glossaulax)
andersoni: 4(1):1-12. Odostomia
(Chesapeakella): 3(1):96. Oenopota
pumilus: 4(1):39-42. Ostrea: 4(1):1-12.
Pachythaerus: 4(2):238. Pelecypoda,
Unspecified: 2:79. Perissitys: 4(2):236.
Plicatula inezana: 4(1):1-12. Pliodon
spp.: 4(1):107. Protothaca: 4(1):1-12.
Purisima: 2:84-85. Pyramidellidae:
3(1):96. Raeta: 4(1):1-12. Rapana
bezoar vaquerosensis, R. imperialis:
2:85-85. Rotella nana: 4(1):39-42.
Sanguinolaria toulai: 4(1):1-12.
Seguenzia, Seguenziacea: 1:92.
Siphocyraea henekeni, Siphonaria
maura pica, Solenosteira: 4(1):1-12.
Spisula confraga, S. modicella:
4(1):39-42. Strombina: 4(1):1-12.
Strombus (Tricornis) costatus, S.
(Tricornis) leidyi, S. (Tricornis) maya-
censis: 4(1):108. Tegu/a spp.:
4(1):1-12. Teinostoma nana:
4(1):39-42. Terebra burckhardti,
Theodoxus, Trachycardium, Trochita
radians, T. spirata, T. trochiformis:
4(1):1-12. Turridae: 3(1):98. Turritella
spp.: 2:84-85. Turritella spp.: 2:84-85;
4(1):1-12. Utriculastra: 4(1):39-42.
Vasum pufferi: 2:84-85. Vermetus
contortus: 4(1):1-12

Palmer, Katherine Van Winkle
Obituary: 1:79-80

Paramyosin
Lasmigona costata: 2:82

Parasitology
Amnicola limosa: 5(1):73-84 (passim).
Ancylus fluviatilis: 3(2):151-168. Biom-
phalaria spp.: 1:67-70, 106; 5(1):85-90.
Bivalvia, Unspecified: 3(1):93. Buli-
nus cernicus, B. forskali Group: 1:07.
B. truncatus: 5(1):85-90. Campeloma
decisum: 5(1):73-84. Caretta caretta:
3(1):93. Corbicula, C. fluminea:
S2:89-94. Crassostrea virginica:
$3:59-70. Fargoa bartschi: $1:1-22
(passim). Ferrissia: 5(1):73-84.
Gastropoda, Unspecified: 3(1):93.
Gyraulus: 2:88. G. parvus, Helisoma
anceps (passim): 5(1):73-84. Leptoxis
carinata: 4(1):119. Lymnaea (Stagni-
cola) elodes: 1:67-70. L. emarginata,
L. stagnalis: 5(1):73-84. Melanoides
tuberculata: 5(1):105-124. Melanopsis:
5(1):85-90. Mellanella spp.: 2:83.
Mercuria confusa, M. punica:
5(1):85-90. Odostomia (Chlysallida):
4(1):122. Onchomelania hupensis:
2:88. Physa integra (passim) 5(1):73-84.

Radix: 2:88. Schistosoma hematobium
1:107. S. japonicum: 2:88. S. mansoni
1:67-70, 106; 4(1):120; 5(1):85-90, S.
mansoni Puerto Rican PR-1, S. man-
soni Puerto Rican PR-2: 1:106.
Sphaerium spp., Tricula: 2:88

Pathology
Aeromonas cCaviae: 2:82. Bankia
gouldi, Boveria teredinidi, B. zeuke-
vitchi, Crassostrea virginica (passim),
Haplosporidium: $1:101-109. H.
costalis: S3:59-70. H. (Minchinia)
nelsoni: S3:17-23, 59-70. Octopus
spp., Pseudomonas stutzeri: 2:93-94.
Teredo spp.: S1:101-109

Penial Complex
Biomphalaria spp., Lymnaea
(Stagnicola) elodes: 1:67-70

Peninsula Effect
Ammonitellidae, Bulimulidae, Haplo-
trematidae, Helminthoglyptidae,
Oreohelicidae, Spiraxidae: 1:97

Periostracum Microstructure
Lithophaga nigra, Pinctada martensi:
1:101

Phenotypes
Goniobasis proxima: 1:105. Nucella
emarginata: 5(1):105-214 (passim)

Phenotype, Shell
See Shell Phenotype

Phosphates
Cionella lubrica: 3(1):27-32

Photoreceptors
Hermissenda crassicornis: 1:13
(passim)

Phylogenetics
Acado: 5(2):215-241. Acanthopleura
granulata: $1:1-22. Aclididae, Aclis,
Acochlidiacea, Acteocina spp.:
Acteocinidae, Acteon: S$1:1-22.
Aeolidacea: 5(2):215-241. Aglaja,
Aglajidae, Akera, Akeridae, Allo-
gastropoda, Amaea, Amphibola,
Amphibolidae, Anaspidea, Angutis-
pira: S1:1-22. Anidolyta, A.
spongotheras: 5(2):215-241. Annelida:
3(2):213-221 (passim). Anthobranchia:
5(2):215-241. Aplacophora: 6(1):57-68.
Aplysia spp., Aplysiidae, Aplysiomor-
pha, Architectonicacea, Architecton-
icidae: S1:1-22. Arminacea:
5(2):215-241. Arthropoda:
3(2):213-221 (passim). Ascoglossa,
Atyidae, Basommatophora: S1:1-22.
Bathyberthella spp.: 5(2):215-241.
Bellamya spp.: 4(1):107. Berthelinia:
$1:1-22. Berthella spp.: 5(2):215-241;
$1:1-22. Berthellina: 5(2):215-241;
$1:1-22. Berthellina citrina, B. engeli,
Berthellinae, Birthellini, Berthellinops:
5(2):215-241. Blauneria, Boonea,
Bulla: $1:1-22. B. membranacea, B.
plumula: 5(2):215-241. Bullidae,

300 AMER

Bullina, Bullomorpha: $1:1-22.
Caelatura: 4(1):107. Calyptraeidae,
Campanile, Campanilidae, Cary-
chium, Cephalaspidea, Cerithiopsa-
cea, Cerithiopsidae: $1:1-22.
Chaetopleura apiculata: 6(1):69-78.
Chelidonura, Chilina, Chilinidae,
Chrysallida: $1:1-22. Cladobranchia,
Cleanthus: 5(2):215-241. Couthou-
yella: S1:1-22. Cyanogaster:
5(2):215-241. Cyclophoridae, Cyclo-
stremella, Cyclostremellidae, Cylin
chna, Cylindrobulla, Cylindrobullidae,
Cymbulia, Cymbuliidae: $1:1-22.
Dendronotacea: 5(2):215-241.
Detracia, Diaphana, Diaphanidae,
Docoglossa: $1:1-22. Doridacea:
5(2):215-241. Ebala, Ellobiidae,
Ellobium, Elysia, Elysiidae, Entomo-
taeniata, Epitoniacea, Epitoniidae,
Epitonium, Eulimacea, Eulimidae:
$1:1-22. Euselenops, E. luniceps:
5(2):215-241. Euthyneura, Fargoa
bartschi: S1:1-22. Gastroplax:
5(2):215-241. Gegania: $1:1-22.
Gigantonotum: 5(2):215-241. Gleba,
Gymnosomata: S1:1-22. Gymnoto-
plax, G. americanus: 5(2):215-241.
Haminoea, Hedylopsidae, Hedylop-
sis, Heliaucus, H. cylindricus, H.
perreieri, Heterobranchia, Hetero-
gastropoda, Heteroglossa,
Hydatina, Hydatinidae, Janthina
spp., uJ. exigua, J. janthina, Jan-
thinidae: $1:1-22. Joannisia:
5(2):215-241. Juliidae: S1:1-22.
Koonsia: 5(2):215-241. Latia, Latiidae,
Leucophytia, Limacinidae, Lima-
pontia, Limapontiidae, Littorina,
Lymacina: $1:1-22. Macfarlandaea:
5(2):215-241. Marinula, Mathilda,
Mathildidae, Maxacteon, Melam-
pidae, Melampus: S1:1-22. Mesodon
zaletus: 2:97-98. Mesogastropoda,
Micromelo, Muricidae, Myxa:
$1:1-22. Neda: 5(2):215-241. Nem-
ertea: 3(2):213-221. Neogastropoda:
$1:1-22. Neopilina: 3(2):213-221.
Neothauma tanganyicense: 4(1):107.
Notaspidea: 5(2):215-241; $1:1-22.
Nudibranchia, Odostomia, Omalo-
gyra: $1:1-22. Ombrella: 5(2):215-241.
Onchidella, Onchidiidae, Onchidium:
$1:1-22. Operculatum: 5(2):215-241.
Opisthobranchia: $1:1-22. Oscani-
opsis, Oscaniella, Oscanius:
5(2):215-241. Otina, Otinidae,
Ovatella, Oxynidae, Oxynoe:
$1:1-22. Parmophorus, Patella
perversa, P umbraculum:
5(2):215-241. Peracle, Peraclidae,
Phanerophthalmus, Philine, Philini-
dae, Philinoglossa, Philinoglossidae,

Philippia, Planorbidae: S1:1-22.
Pleurehdera, P. haraldi: 5(2):215-241.
Pleurobranchacea, Pleurobranchaea
spp., Pleurobranchaeidae, Pleuro-
branchella spp.: 5(2):215-241.
Pleurobranchidae: 5(2):215-241;
$1:1-22. Pleurobranchidium, Pleuor-
branchillus, Pleurobranchinae,
Pleurobranchoides gilchristi:
5(2):215-241. Pleurobranchomorpha:
$1:1-22. Pleurobranchus:
5(2):215-241; S1:1-22. Pleurobranchus
spp.: 5(2):215-241, 243-258. Pliodon
ovata, P spekil: 4(1):107. Polyplaco-
phora: 6(1):57-68. Prosobranchia:
$1:1-22. Protostomia: 3(2):213-221
(passim). Pseudomalaxis, Pseudo-
skenella, Ptenoglossa, Pulmonata,
Pupa, Purpura patula, Pyramidella
crenulata, Pyramidellacea, Pyrami-
dellidae, Pythia, Radix, Retusidae,
Retussa: $1:1-22. Rhinocoela:
3(2):213-221 (passim). Ringicula,
Ringiculidae, Rissoella, Rissoellidae,
Roxania: $1:1-22. Roya, R.
spongotheras: 5(2):215-241.
Sacoglossa, Salinator, Sayella,
Scaphander, Scaphandridae: S1:1-22.
Siphonaria: 5(2):215-241; $1:1-22.
Siphonariidae, Smaragdinella:
$1:1-22. Spiricella: 5(2):215-241.
Stiligar, Stiligeridae: S1:1-22.
Susania: 5(2):215-241. Systellom-
matophor, Thecosomata, Toledella:
$1:1-22. Triodopsis: 2:97-98.
Triopohridae, Trochidae, Turbonilla, T.
vineae, Turritellidae: $1:1-22.
Tylodina: 5(2):215-241. T. alfredensis:
5(2):243-258. T. spp., Tylodinella, T.
trinchesii, Tylodinidae, Umbraculacea:
5(2):215-241. Umbraculidae, Um-
braculum: 5(2):215-241; $1:1-22. U.
umbraculum, Umbrella: 5(2):215-241.
Valvata, Valvatacea, Valvatidae,
Veronicellidae: S1:1-22. Viviparidae:
3(1):107. Volvatella, Volvatellidae:
$1:1-22. Williamia: 5(2):215-241;
$1:1-22

Physiology

Aciculidae: 3(2):223-231. Amoeba
proteus: S1:79-83. Ampullariidae:
3(2):223-231. Ancylus fluviatilis:
3(2):135-142, 151-168, 243-265,
269-272. Anodonta grandis:
3(2):233-242. Aplysiopsis zebra:
5(2):259-280. Argopecten irradians:
$1:59-78. Ascobulla ulla:
5(2):259-280. Assimineidae:
3(2):223-231. Australorbis glabratus:
3(2):213-221. Berthelinia caribbea:
5(2):259-280. Biomphalaria glabrata:
3(2):213-221; S1:79-83. Bithynia:
3(2):135-142 (passim), 269-272. B.

. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

tentaculata: 3(2):179-186. Bithyniidae:
3(2):223-231. Bittum alternatum:
$1:85-91. Bosellia mimetica, Boselli-
dae: 5(2):259-280. Buccinum un-
datum: 3(2):223-231. Caliphyllidae:

5(2):259-280. Carunculina texasensis:

3(2):233-242. Cerithiidae:
3(2):223-231. Chaetomorpha:
5(2):259-280. Colpidium: S$1:79-83.
Corbicula fluminea: 3(1):101;
3(2):233-242, 267-268, 269, 272;

4(1):81-88. Corbiculacea: 3(2):201-212.

Costasiella ocellifera, C. nonatoi,
Costasiellidae: 5(2):259-280. Crasso-
strea virginica: S1:79-83; S3:41-49.
Crepidula fornicata: 3(2):135-142
(passim); S1:85-91. C. plana:
$1:85-91. Cryptozona belangeri:
4(1):114; 4(2):237. Cyclophoridae:
3(2):223-231. Cyerce antillensis:
5(2):259-280. Cymatium parthen-
opeum: S1:85-91. Daphnia: $1:79-83.
Deroceras reticulatum: 3(2):223-231.
Elimia potosiensis: 3(1):100. Elysia
spp., Elysiidae, Ercolania funerea, E.
fuscata: 5(2):259-280. Ferrissia
rivularis: 3(2):135-142 (passim),
243-265. F. wautieri: 3(2):151-168.
Fusconaia ebena: 5(2):177-179.
Haliotis corrugata, H. rufescens,
Helicinidae: 3(2):223-231. Helisoma:
$1:51-58. H. anceps: 4(1):118-119. H.
trivolvis: 3(2):213-221, 243-265;
4(1):118-119. Helix pomatia:
3(2):223-231. Hiatella: 3(2):135-142

(passim). Hydrobiidae, Hydrocenidae:

3(2):223231. Illex illecebrosus:
3(1):107; 4(1):55-50. Laevapex fuscus:
3(2):243-265 (passim). Lampsilis
claibornensis: 3(2):233-242.
Lasmigona costata: 2:35-40. Leptoxis
arkansensis: 3(1):100. L. carinata:
3(2):169-177, 269-272. Ligumia
subrostrata: 3(2):233-242; 5(1):41-48;
$1:51-58. Limapontia capitata:
5(2):259-280. Limax pseudoflavus:
3(2):223-231. Liolophura gaimardi:
$1:79-83. Littorina irrorata:
3(2):223-231. L. littorea: 3(2):135-142
(passim). Lobiger souverbiei:
5(2):259-280. Loligo forbesi: 4(2):240.
Lymnaea (Stagnicola) elodes:
3(2):143-150, 213-221, 269-272. L.
palustris: 3(2):213-221. L. peregra:
3(2):135-142 (passim). L. stagnalis:
3(2):135-142 (passim), 223-231;
$1:51-58. Macoma balthica:
3(2):213-221. Margaritifera hembeli:
3(2):233-242. Marisa cornuarietis:
3(2):223-231. Melampus bidentatus:
3(2):135-142 (passim); 4(1):110-111;
4(2):236-237. Melaniidae, Melanopo-
sidae, Mesogastropoda: 3(2):223-231.

AMER. MALAC. BULL. SUBJECT INDEX

Mollusca, Unspecified: 3(2):135-142
(passim). Monas: S1:79-83. Mopalia
mucosa: S2:85-91. Mourgona ger-
maineae: 5(2):259-280. Musculium:
3(2):269-272. M. lacustre: 3(2):187-200.
M. partumeium: 3(2):187-200, 201-212.
M. securis: 3(2):187-200. Mya
arenaria: S1:79-83. Mytilus edulis:
3(1):33-40; 3(2):213-221; S1:79-83,
85-91. Nautilus: S1:51-58. Nerita
fulgurans, Neritacea, Neritidae,
Neritina latissima: 3(2):223-231. Oc-
topus dolfleini: 2:91. O. vulgaris:
4(2):240. Oxynoe antillarum, O.
azuropunctata: 5(2):259-280. Patella
aspersa: 3(1):33-40. P vulgata:
3(1):33-40; 3(2):223-231. Periplaneta
americana: $1:79-83. Physa fontinalis:
3(2):135-142 (passim), 243-265.
Physella virgata: 3(2):269-272. P
virgata virgata: 3(2):243-265.
Pisidiidae: 3(2):201-212. Pisidium:
3(2):269-272. P spp.: 3(2):187-212;
5(1):41-48. Placida dendritica, P
kingstoni: 5(2):259-280. Planorbis
corneus: 3(2):135-142 (passim),
213-221. Pleurocera acuta: 3(1):100.
Pleuroceridae: 3(2):223-231. Polinices
duplicatus: 3(2):135-142 (passim).
Pomacea lineata: 3(2):223-231. P
paludosa: S$1:51-58. Potamopyrgus
jenkinsii: 3(2):223-231. Rangia
cuneata: 3(2):233-242. Rissoacea,
Rissoidae: 3(2):223-231. Schistom-
soma mansoni: $1:79-83. Sepia of-
finalis: 4(2):240. Sphaerium spp.:
3(2):187-200, 201-212; 5(1):41-48.
Spirodon carinata: 3(2):169-177.
Spisula solidissima: 3(2):135-142
(passim). Stiligeridae: 5(2):259-280.
Strombus gigas, Syrnolopsidae,
Thiaridae: 3(2):223-231. Unio pic-
torum: 3(2):233-242. Unionacea:
3(2):201-212. Valvata piscinalis:
3(2):243-265. Valvatacea, Valvatidae,
Viviparacea, Viviparidae: 3(2):223-231.
Viviparus: 3(2):269-272. V. spp.:
3(2):223-231. Volvatella bermudae:
5(2):259-280

Physiology, Comparative
Cardium edule, Crepidula spp.,
Gelonia erosa, Geukensia demissa,
Modiolus demissa, Modiolus modio-
lus, Mytilus californianus: 3(1):33-40

Pigment Patterns
Mollusca, Unspecified: 4(2):242

Plant Associations, freshwater
Unionidae: 1:61-68

Poecilogony
Acteonia sp., A. candei, A. lepta,
Tornatina spp.: 3(1):98

Population Dynamics
Corbicula fluminea: S2:89-94

Population History
Cepaea sp.: 1:103

Population Structure
Crassostrea virginica: 1:108

Predation
Alvania auberiana: 4(2):185-199.
Ancipenser transmontanus: S2:7-39.
Anemonia sulcata: 5(2):185-196.
Aplocinotus grunniens: S2:7-39,
89-94. Argopecten arquisulcatus:
4(2):241-242. Ascophyllum: 1:92.
Asterias amurensis: 2:94. A. forbesi:
$3:59-70. Aythya affinis, A. marila:
$3:59-70. Berryteuthis anonychus:
4(2):241. Bittum varium: 4(2):185-199.
Boonea, B. impressa, Busycon sp.,
B. canaliculatum, Callinectes
sapidus: S3:59-70. Cambarus bartonii:
$2:89-94, 211-218. Carcinus maenas:
4(1):108. Collisella pelta: 2:80. Cor-
bicula fluminea: S2:7-39, 89-94,
211-218. Corphium, Crassostrea vir-
ginica: S3:59-70. Crossater papposis:
5(2):287-292. Crucibulum spinosum:
4(2):241-242. Cyprinus carpio:
$2:89-94. Dugesia tigrina: S2:7-39,
89-94. Epitonium albidum: 1:1-12.
Eupleura caudata, Eurypanopeus
depressus: S3:59-70. Favorinus
branchialis: 5(2):185-196. Fundulus:
2:1-20. Gonatus middendorfi:
4(2):241. Graptemys pulchra: S2:7-39.
Haemopsis grandis: 5(1):73-84.
Haliotis cracherodii: 4(2):233-234.
Halisarca dujardini: 4(1):103-104. Ic-
talurus furcatus: S2:7-39, 89-94. /.
punctatus: S2:89-94, 211-218. Ictiobus
bubalus: S2:7-39, 89-94. |. cyprinellus:
$2:7-39. |. niger: S2:7-39, 89-94. Illex
illecebrosus: 1:90. Laevicardium
substriatum: 4(2):241-242. Leiostomus
xanthurus: S3:59-70. Lepomis gib-
bosus: 5(1):73-84. L. microchirus:
$2:89-94. L. microlophus: 5(1):73-84;
$2:7-39, 89-94. Limulus polyphemus:
2:96. Littorina filosa: 4(1):112. L. lit-
torea: 1:92. L. obtusata: 1:92;
4(1):108. L. scabra: 4(1):112. Lymnaea
(Stagnicola) elodes: 5(1):73-84.
Marginella aureocincta: 4(2):185-199.
Megalodonta beckil: 5(1):73-84.
Melanitta fusca, M. nigra: S3:59-70.
Metopograpsus: 4(1):112. Metridium
senile: 5(2):287-292. Micropogon un-
dulatus: S3:59-70. Minytrema
melanops: S2:7-39, 89-94. Mitra idae:
1:91-92. Molgula manhattensis:
$3:59-70. Moroteuthis pacifica, M.
robusta: 4(2):241. Mytilus edulis:
2:63-73. Navanax inermis:
5(2):287-292. Neopanope Sayi:
$3:59-70. Nucella lapillus: 1:92.
Nudibranchia: 5(2):287-292. Octopus

: 1983 - 1988 301

spp.: 4(2):233-234. O. bimaculoides:
4(2):241-242. Odostomia: S3:59-70.
Ommastrephes bartrami: 4(2):241.
Ospanus tau: S3:59-70. Orconectes
spp.: 5(1):73-84; S2:211-218. Ostrea
equestris: 2:63-73. Pachygrapsus
crassipes: 2:1-20. Panopeus herbstii:
2:1-20, S3:59-70. Perkinsus marinus:
$3:59-70. Phascolosoma agassizii:
1:91-92. Pogonias cromis: S3:59-70.
Pleurobranchaea californica:
5(2):287-292. Potamogeton:
5(1):73-84. Procambarus clarkii:
$2:89-94, 211-218. Procladius culici-
formis: S2:7-39. Procyon lotor:
$2:7-39, 89-94. Promenetus exacuous:
5(1):73-84. Pseudopleuronectes
americanus: 5(2):287-292. Rallus
crepitans: 2:1-20. Rangia cuneata:
2:63-73. Rhinoptera bonasus, Rithro-
panopeus harrisii: S3:59-70. Rossia
pacifica: 2:91-92. Salmo trutta:
5(1):73-84. Saxidomus nuttalli,
Semele decisa: 4(2):241-242.
Squalus: 2:91-92. Stichodactyla
helianthus: 1:1-12. Stylochus, S. ellip-
ticus: S3:59-70. Tautogolabris
adspersus: 5(2):287-292. Thais
deltoidea: Thalamita crenata: 4(1):112.
Thalassoma bifasciatum: 1:8. Theba
pisana: 1:104. Thunnus alalunga:
4(2):241. Umbra limi: 5(1):73-84.
Urosalpinx cinerea: S3:59-70.
Vallisneria americana: 5(1):73-84
Preservation
Loligo sanpaulensis: 6(2):213-217
Proboscis
Janthina sp.: 1:4, 7, 9, 10. Thais
haemastoma canaliculata: 2:63-73
Proton Probe Analysis
Crassostrea rhizophorae: 1:102
Radiotracers
Corbicula fluminea: S2:219-222
Radula
Acanthopleura, A. granulata:
4(1):114-115. Acmaeidae: 4(1):115.
Adalaria lovéni, A. pacifica, A. prox-
ima: 2:95. Ancylus fluviatilis:
3(2):151-168. Aplacophora: 6(1):57-68.
Aplysiopsis zebra, Ascobulla ulla,
Berthelinia caribbea, Bosellia mimet-
ica, Bosellidae: 5(2):259-280. Buc-
cinanops: 3(1):101-102. Buchanania,
B. onchidioides: 2:21-34. Bullia:
3(1):101-102. Caliphyllidae:
5(2):259-280. Cellana: 4(1):115.
Cerithidea spp., Cerithideopsilla,
Cerithideopsis: 2:1-20. Chaetomorpha:
5(2):259-280. Chiton, Chitonidae:
4(1):114-115. Clypeomorus spp.:
4(1):109. Costasiella ocellifera, C. non-
atoi, Costasiellidae, Cyerce antillensis:
5(2):259-280. Dondersiidae: 6(1):57-68.

302 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Elysia spp., Ercolania funerea, E.
fuscata: 5(2):259-280. Fissurellidea
spp.: 2:21-34. Fusinus acanthodes, F.
(Pagodula) acanthodes, Fusus acanth-
odes: 3(1):101-102. Gastropoda,
Unspecified: 4(2):233, 244. Graptacme
calamus: 1:100. Lepetidae: 4(1):115.
Limapontia capitata, Lobiger souver-
biei: 5(2):259-280. Lyria guidingii:
3(1):101-102. Mancinella, M. alouina:
4(1):110. Mourgona germaineae:
5(2):259-280. Nassariidae:
3(1):101-102. Nucella: 4(1):110. N.
lapillus: 4(1):110. Onchidoris spp.:
2:95. Oxynoe antillarum, O. azuro-
punctata: 5(2):259-280. (Pagodula):
3(1):101-102. Patella, Patellidae,
Patellogastropoda: 4(1):115. Physa:
6(1):57-68. Placida dendritica, P
kingstoni: 5(2):259-280. Pleurotomaria
atlantica: 3(1):101-102. Polycarpa,
Polyplacophora: 6(1):57-68.
Pupillaea, P annulus, P aperta:
2:21-34. Purpura: 3(1):101-102;
4(1):110. P persica, Purpurella, P
patula: 4(1):110. Rhodope: 6(1):57-68.
Seguenziacea: 1:92. Simrothiella:
sp.: 6(1):57-68. Solariella carvalhoi:
3(1):101-102. Stiligeridae: 5(2):259-280.
Thais: 4(1):110. T haemostoma
canaliculata: 2:63-73. T. nodosa:
4(1):110. T. nodosa mevetricula,
Trophon acanthodes, T. (Pagodula)
acanthodes, Typhina riosi, Volutidae:
3(1):101-102. Volvatella bermudae:
5(2):259-280

Recruitment
Amblema plicata, Fusconaia ebena:
6(1):49-54. Periploma margaritaceum:
2:35-40

Regeneration
Thais haemastoma canaliculata pro-
boscis: 2:63-73. Tegula sp. shell:
1:102

Reproduction
Actinonaias ellipsiformis: 3(1):93.
Adalaria, A. proxima, Aeolidia papil-
losa: 5(2):293-301. Alloteuthis:
4(2):217-227. Ancylus fluviatilis:
3(2):151-168. Anguispira alternata:
4(2):237. Anodonta spp.: 3(1):93;
5(1):91-99. Anodontoides ferussaci-
anus: 3(1):93. Argonauta: 4(2):217-227.
Ashmunella levettei, A. varicifera:
1:21-26. Bathypolypus arcticus:
4(2):217-227. Batissa (Cyrenobatissa)
subsulcata: 5(1):91-99. Bulinus
tropicus: 1:96. Corbicula fluminalis:
5(1):91-99. C. fluminea: 5(1):91-99;
$2:99-111, 133-142, 193-201, 211-218.
Crassostrea virginica: S3:25-29,
41-49. Eledone cirrhosa, E. moschata,
Eledonella pygmaea: 4(2):217-227.

Epitonium spp.: 1:1-12. Euprymna:
4(2):217-227. Fusconaia flava: 3(1):93.
Idiosepius, Illex: 4(2):217-227. |. ille-
cebrosus: 4(2):239. Lamprotula leai:
5(1):91-99. Lampsilis ovata, L. radiata,
Lasmigona compressa: 3(1):93. Lim-
noperna fortuei: 5(1):91-99. Loligo
vulgaris: 4(2):217-227. Lymnaea
(Stagnicola) elodes: 3(2):143-150.
Melampus bidentatus: 4(1):121-122.
Musculium lacustre: 5(1):91-99.
Nassarius pauperatus: 5(2):293-301
(passim). Nautilus, Octopodidae, Oc-
topus spp., O. briareus: 4(2):217-227.
O. burryi: 2:92. O. vulgaris:
4(2):217-227. Onchidoris spp.:
5(2):293-301. Orbicularia: 5(2):159-164
(passim). Ostrea edulis, O. lurida:
4(1):61-79 (all passim). Phestilla
sibogae: 5(2):293-301 (passim).
Pisidiidae: 3(1):100; 4(1):61-79.
Pisidium annandalei, P_ clarkeanum:
5(1):91-99. Planaxidae, Planaxis:
3(1):96. Polymesoda (Geloina) erosa:
5(1):91-99. Pteroctopus tetracirrhus,
Rossia, Sepia spp., Sepietta,
Sepiola, Spirula: 4(2):217-227. Thais:
5(2):293-301 (passim). Tremoctopus:
4(2):217-227. Union douglasiae:
5(1):91-99. Unionidae, Unspecified:
4(1):61-79. Vampyroteuthis, V. infer-
nalis: 4(2):217-227. Viviparus
georgianus: 3(2):268

Salinity
Bankia gouldi: 4(1
strea virginica: 4(1
schi, T. navalis: 4(

Sampling Methods
Unionids, unspecified: 1:93

Sensory Hairs
Polyplacophora: 6(1):141-151

Sensory Organs
Acanthochiton fascicularis, Chaeto-
pleura lurida, C. peruviana:
6(1):141-151. Chiton olivaceus:
6(1):131-139, 141-151. Corbicula
fluminea: 1:13-20. Donax trunculus:
1:13 (passim). Eudoxochiton nobilis,
Ischnochiton herdmani, Katharina
tunicata, Lepidochitona cinerea, L.
dentiens, Lepidozona retiporosus:
6(1):141-151. Lepidopleurus cajetanus:
6(1):141-151, 153-159. Mopalia spp.,
Placiphorella velata, Plaxiphora
obtecta, Tonicella insignis: 6(1):141-151

Sexual Dimorphism
Aforia circinata: 2:82. Elliptio icterina:
1:95. E. lanceolata: 1:94-95. Villosa
villosa: 1:95

Sexuality
Bankia, Calyptraeidae, Corbicula,
Crassostrea virginica, Crepidula,
Epitonium albidum, Mercenaria,

189-99. Crasso-
1101. Teredo bart-

)
)
1):89-99

Ostrea gigas, Teredinidae, Teredo
navalis: 3(1):85-88

Shallow Water Marine Fauna
Paleontology: 2:79-80

Shell
Conus: 3(1):95. Gastropoda,
Unspecified: 2:80-81; 3(1):95. Lissarca
notocadensis: 4(2):235. Lottia
gigantea: 4(2):242-243. Mytilus edulis:
2:41-50

Shell Ashing
Lasmigona subviridis, Medionidus
conradicus, Pleurobema oviforme,
Villosa vanuxemi: 6(2):179-188

Shell Calcium
Ancylus fluviatilis, Biomphalaria
glabrata, B. pfeifferi, Cincinnatia cin-
cinnatiensis (passim), Fedrrissia
rivularis (passim), Helisoma anceps
(passim), Lymnaea (Stagnicola)
elodes, L. peregra (passim), Nucella
lapillus (passim), Physella gyrina
(passim), P. integra (passim):
5(1):105-124. Pinctada martensi: 1:101.
Planorbis corneus, Sphaerium spp.:
5(1):105-124 (all passim). Thais
haemastoma canaliculata:
6(2):189-197. Valvata tricarinata:
5(1):105-124 (passim)

Shell Chemistry, Minor Elements
Crassostrea gigas, C. rhizophorae,
Ostrea lurida: 1:102

Shell Chemistry, Trace Elements
Crassostrea gigas, C. rhizophorae,
Ostrea lurida: 1:102

Shell Color Patterns
Cepaea nemoralis: 3(1):1-10. C.
nemoralis nemoralis, C. vindobonensis:
1:107-108. Corbicula fluminea: 2:87.
Liguus fasciatus: 1:98; L. spp.:
3(1):1-20. Littorina saxatilis: 3(1):1-10.
Nucella emarginata, Thais emarginata:
1:105. Theba pisana: 1:104, 104-105

Shell Formation
Aeolidia papillosa: 6(1):57-68.
Amblema costata, Anodonta grandis:
$1:35-50. Chiton polii, Epimenia ver-
rucosa, Halomenia gravida: 6(1):57-68.
Helisoma duryi, Helix pomacea:
$1:35-50. /schnochiton rissoi, Lepi-
dochitona cinerea, L. corrugata:
6(1):57-68. Lymnaea stagnalis, Mer-
cenaria mercenaria: $1:35-50. Mid-
dendorffia caprearum: 6(1):57-68.
Mytilus edulis: S1:35-50.
Nematomenia banyulensis, N. protec-
ta, Neomenia carinata, Neopilina:
6(1):35-50. Samarangia quadrangu-
laris: S1:35-50. Septemchiton:
6(1):35-50

Shell Microstructure
Calyptogena magnifica: 1:101. Cor-
bicula fluminea: 2:87; 3(1):100-101;

AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988 303

4(1):116-117; 4(2):234. Crassatella
ponderosa, C. vadosa, Crassatelli-
dae: 4(2):238. Crassostrea rhizo-
phorae: 1:35-42. Geukensia demissa
demissa: 5(1):173-176. G. demissa
granosissima: 3(1):103; 4(1):112;
5(1):173-176. Lyonsia californica:
5(1):173-176 (passim). L. floridana:
2:41-50. Mytilidae: 1:101. Mytilimeria
nutalli: 5(1):173-176 (passim).
Pachythaerus: 4(2):238. Pinctada
martensi: 5(1):173-176 (passim).
Polymesoda caroliniana: 4(1):116-117;
4(2):234; 6(2):199-206. Tegula sp.:
1:102; 2:41-50. Vesicomyidae: 1:101

Shell Mineralization
Argopecten irradians, Helisoma,
Ligumia subrostrata, Lymnaea
stagnalis, Nautilus, Pomacea
paludosa: S$1:51-58. Thais haemo-
stoma canaliculata: 6(2):189-197

Shell Morphology
Acanthochiles hemphilli, Acantho-
chitona spp.: 6(1):79-114, 115-130.
Acanthopleura vaillantii: 6(1):115-130.
Aforia circinata: 2:82. Aligena cokeri:
1:91. Arca noae, Arctica islandica,
Argopecten irradians: S1:59-78. Ash-
munella lenticula, A. proxima albi-
caudata: 1:106. Astarte castanea:
$1:59-78. Brachidontes exustus:
4(2):233-234. Callistochiton adenensis:
6(1):115-130. Campeloma geniculum,
C. parthenum: 3(1):99. Chiton spp.:
6(1):115-130. Choreplax lata:
6(1):79-114. Crassostrea virginica:
$1:59-78. Cryptoconchus floridanus:
6(1):79-114. Cyclocardia borealis, En-
sis directus: S1:59-78. Geukensia
demissa: 4(2):233-234; $1:59-78.
Graptacme calamus: 1:100. lo
fluvialis: 5(1):65-72 (passim). Ischno-
chiton winckworthi, |. yerburyi:
6(1):115-130. Lepidochitona dentiens:
4(2):243. Lepidozona luzonica:
6(1):115-130. Macoma balthica:
$1:59-78. Mancinella, M. alouina:
4(1):110. Mercenaria mercenaria,
Modiolus modiolus, Mulinia lateralis,
Mya arenaria, Mytilus californianus,
M. edulis: S1:59-78. Nucella, N.
lapillus: 4(1):110. Notoplax (Notoplax)
arabica, Onithochiton erythraeus:
6(1):115-130. Pholadidae, Placopecten
magellanicus: S1:59-78. Pupilla spp.:
1:99. Purpura, P. persica, Purpurella,
P. patula: 4(1):110. Spisula solidissima:
$1:59-78. Thais, T. nodosa: 4(1):110.
Tonicia (Lucilina) sueziensis:
6(1):115-130

Shell Phenotypes
Partula spp.: 1:103-104. Theba
pisana: 1:104, 104-105

Shell Protein
Crassostrea virginica: 2:41-50

Shell Secretion
Mytilidae: 1:101. Tegula sp.: 1:102.
Vesicomyidae: 1:101

Shell Structure
Mercenaria: 3(1):85-88

Siphons
Bankivia: 3(1):95. Cardiomya
planetica: 1:13. Corbicula fluminea:
1:13-20. Gastropoda, Unspecified,
Trochidae, Turritellidae, Umbonium,
Vermetidae: 3(1):95

Site Transplantation
Arianta arbustorum, Cepaea hortensis:
1:103. Polymesoda caroliniana:
6(2):199-206

Size
Mercenaria mercenaria: 1:107

Spawning
Batillaria minima, Cerithidea spp.:
2:1-20. Corbicula fluminea: 4(1):116;
S2:69-81. Periploma margaritaceum:
2:35-40

Speciation
Ammonitellidae, Bulimulidae, Haplo-
trematidae, Helminthoglyptidae: 1:97.
Melanoides tuberculata: 5(1):105-124
(passim). Oreohelicidae, Spiraxidae:
1:97

Spermatheca Ultramorphology
Biomphalaria glabrata: 1:96-97

Spermatophore
Batillaria minima, Cerithidea scalari-
formis: 2:1-20

Starvation
Thais: 3(2):213-221 (passim)

Statocyst
Helix vulgaris: 1:13 (passim). Lolli-
guncula brevis: 1:90. Lymnaea
stagnalis: 1:13 (passim)

Statolith
Illex illecebrosus: 2:51-56;
4(2):240-241. Loligo opalescens: 2:93

Swimming
Aplysia brasiliana, Aplysia califor-
nica: 2:78

Taxonomy
Acado: 5(2):215-241. Acanthochiles
hemphilli, Acanthochitona spp.:
6(1):79-114, 115-130. Acanthodoris:
5(2):243-258. Acanthopleura vaillantii:
6(1):115-130. Acteocina smithi, Acteon
flammeus, A. fortis, Acteonidae:
5(2):243-258. Actinonaias spp.:
6(1):19-37. Adamete viridula,
Admetula, A. evulsa: 2:57-61.
Aegries: 5(2):243-258. Aeolidacea:
5(2):215-241. Aeolidiella alba, A. in-
dica, Aeolidiidae, Aglajidae, Akera
soluta, Akeridae: 5(2):243-258.
Alasmidonta spp.: 6(1):19-37. Aldisa
benguela, A. trimaculata, Aldisidae,

Amanda armata: 5(2):243-258.
Amblema spp.: 6(1):19-37. Ampulla
Purpurea: 2:57-61. Anaspidea, Ancula,
Anaspidea: 5(2):243-258. Anidolyta,
A. spongotheras: 5(2):215-241.
Anisodoris prea: 5(2):183-184.
Anodonta spp.: 5(1):91-99; 6(1):19-37.
Anoaontoides ferrussacianus:
6(1):19-37. Anthobranchia: 5(2):215-241.
Aphelodoris brunnea, Aplysia dacty-
lomela, Aplysia spp., Aplysiidae,
Aplysiopsis sinusmensalis:
5(2):243-258. Arcidens confragosus:
6(1):19-37. Armina gilchristi:
5(2):243-258. Arminacea: 5(2):215-241.
Arminidae, Artachaea, Arthritica
hulmei, Ascobulla fischeri, Asterono-
tidae, Atagema gibba, A. rugosa,
Atys cylindrica, Baeolidida palythoae:
5(2):243-258. Bathyberthella, Bathy-
berthella antarctica, B. zelandiae:
5(2):215-241. Bathydorididae:
5(2):243-258. Batissa (Cyrenobatissa)
subsulcata: 5(1):91-99. Berthelinia
schlumbergeri: 5(2):243-258. Berth-
ella spp.: 5(2):215-241, 243-258. Ber-
thellina, B. citrina, B. engeli, Berthel-
linae, Birthellini, Berthellinops:
5(2):215-241. Bivetiella: 2:57-61.
Bonsia nakaza, Bornella anguilla, B.
Stellifer, Bornellidae: 5(2):243-258.
Buccinum spp.: 2:57-61. Bulla am-
pulla: 5(2):243-258. B. membranacea,
B. plumula: 5(2):215-241. Bullidae,
Bullina lineata, Bullinidae, Bursatella
leachii africana, B. leachii leachii,
Cadlina, Caliphyliidae: 5(2):243-258.
Callistochiton adenensis: 6(1):115-130.
Caloria, C. indica: 5(2):243-258.
Cancellaria spp., Cancellariidae:
2:57-61. Carunculina spp.: 6(1):19-37.
Catriona casha: 5(2):243-258. C.
maua: 5(2):183-184. Cephalaspidea,
Ceratophyllidia africana, Ceratosoma,
C. cornigerum, Chelidoneura fulvi-
punctata, C. hirudinina: 5(2):243-258.
Chiton spp.: 6(1):115-130. Choneplax
lata: 6(1):79-114. Chromodorididae,
Chromodoris spp.: 5(2):243-258.
Cladobranchia, Cleanthus:
5(2):215-241. Conradilla caelata:
6(1):19-37. Corambe, Corambidae:
5(2):243-258. Corbicula spp.:
$2:7-39, 113-124. C. fluminalis:
5(1):91-99; S2:113-124. C. fluminea:
4(1):81-88; 5(1):91-99; S2:7-39,
113-124. C. leana: 4(1):81-88; S2-39.
C. manilensis: 4(1):81-88; S2:7-39.
Cratena capensis, C. simba, Craten-
idae, Crimora: 5(2):243-258. Cryp-
toconchus floridana: 6(1):79-114.
Cumberlandia monodonta: 6(1):19-37.
Cuthona spp.: 5(2):243-258.

Cyanogaster: 5(2):215-241.
Cyclonaias spp.: 6(1):19-37. Cylichna
tubulosa, Cylindrobullidae:
5(2):243-258. Cyprogenia irrorata, C.
stegaria: 6(1):19-37. Daphne:
5(2):183-184. Delphinula
trigonostoma: 2:57-61. Den-
drodorididae, Dendrodoris spp.:
5(2):243-258. Dendronotacea:
5(2):215-241. Dermatobranchus,
Discodorididae, Discodoris fragilis:
5(2):243-258. Dondice: 5(2):183-184.
Dolabella auricularia, Dolabrifera
dolabrifera: 5(2):243-258. Doridacea:
5(2):215-241, 243-258. Dorididae,
Doriodoxa benthalis, Doriopsilla, D.
miniata, Doriopsis pecten, Doris ver-
rucosa, Doto spp., Dotoidae:
5(2):243-258. Dromus spp.:
6(1):19-37. Durvilledoris leminiscata:
5(2):243-258. Dysnomia arcaeformis:
6(1):19-37. D. spp., Ellipsaria line-
olata, Elliptio spp.: 6(1):19-37. E.
dilatatus delicatus: 5(2):165-171.
Elysia spp. Elysiidae: 4(2):232;
5(2):243-258. Embletonia gracilis,
Embletoniidae, Endodontidae:
5(2):243-258. Epioblasma spp.:

6(1):19-37. Eubranchidae, Eubranchus:
5(2):243-258. E. coniclus: 5(2):183-184.
Euselenops: 5(2):215-241. E. luniceps:

5(2):215-241, 243-258. Facelina oliva-
cea, Facilinidae, Favorinus ghanensis,
F. japonicus: 5(2):243-258. Fiona pin-
nata, Fionidae, Flabellina, F. capen-
sis, F. funeka, Flabellinidae:
5(2):243-258. Fusconaia spp.:
6(1):19-37. Garamella: 5(2):243-258.
Gastroplax: 5(2):215-241. Gastropter-
idae, Gastropteron alboaruantium, G.
flavobrunneum, Geitodoris capensis:
5(2):243-258. Gigantonotum:
5(2):215-241. Glaucidae, Glaucus
atlanticus, Glossodoris atromarginata,
G. sp., Godiva quadricolor,
Goniodoridae, Goniodoris mercurialis,
G. ovata: 5(2):243-258. Gulo:
5(2):183-184. Gymnodorididae, Gym-
nodoris spp.: 5(2):243-258. Gymno-
toplax, G. americanus: 5(2):215-241.
Halgerda spp.: 5(2):243-258. Hallaxa
apefae: 5(2):183-184. Haminoea
alfredensis, H. natalensis, Haminoei-
dae: 5(2):243-258. Hemistena lata:
6(1):19-37. Hexabranchidae, Hexa-
branchus sanguineus, Hydatina spp.,
Hydatinidae, Hypselodoris spp.:
5(2):243-258. Ischnochiton winck-
worthi, |. yerburyi: 6(1):115-130.
Janolidae, Janolus capensis, J.
longidentatus: 5(2):243-258. Joan-
nisia: 5(2):215-241. Jorunna tormen-
tosa, J. zania, Julia zebra, Kalinga

ornata, Kaloplocamus ramosus, Ken-
trodorididae: 5(2):243-258. Koonsia:
5(2):215-241. Lamprotula leai:
5(1):91-99. Lampsilis spp., Lasmigona
spp., Lastena lata: 6(1):19-37.
Lecithophorus capensis, Leminda
millecra, Lemindidae: 5(2):243-258.
Lemiox rimosus: 6(1):19-37. Lepi-
dozona luzonica: 6(1):115-130. Lep-
todea fragilis, L. leptodon, Lexingtonia
dolabelloides, L. dolabelloides con-
radi, Ligumia recta latissima, L.
subrostrata: 6(1):19-37. Limacia
clavigera: 5(2):243-258. Limnoperna
fortuei: 5(1):91-99. Lobiger souverbiei,
Lophopleurella capensis:
5(2):243-258. Macfarlandaea:
5(2):215-241. Marianina rosea,
Marianinidae, Marioniopsis cyano-
branchiata: 5(2):243-258. Medionidus
conradicus, Megalonaias gigantea,
M. nervosa: 6(1):19-37.
Melanoclamys, Melibe spp.,
Micromelo undata: 5(2):243-258.
Miesea: 5(2):183-184. Mollusca,
Unspecified: 3(1):107. Mordilla brockii:
5(2):243-258. Murex spp.: 2:57-61.
Musculium lacustre: 5(1):91-99.
Nassarius: 2:57-71. Neda:
5(2):215-241. Nembrotha lineolata, N.
livingstonei, Neocorbicula, Notarch-
idae: 5(2):243-258. Notaspidea:
5(2):215-241, 243-258. Notobryon
wardi: 5(2):243-258. Notoplax
(Notoplax) arabica: 6(1):115-130.
Noumea spp., Nudibranchia:
5(2):243-258. Obliquaria reflexa,
Obovaria spp.: 6(1):19-37. Okadaia
elegans, Okenia mediterranea:
5(2):243-258. Ombrella: 5(2):215-241.
Onchidorididae: 5(2):243-258.
Onithochiton erythraeus: 6(1):115-130.
Operculatum, Oscaniopsis, Oscani-
ella, Oscanius: 5(2):215-241. Oxynoe
viridis, Oxynoidae: 5(2):243-258. Par-
mophorus, Patella perversa, P um-
braculum: 5(2):215-241. Pegia fabula:
6(1):19-37. Phanerophthalmus
smaragdius: 5(2):243-258. Phestilla
lugubris: 5(2):185-186. P. melano-
branchia, Philinopsis capensis, P.
cyanea, Phyllida, P varicosa, Phyl-
lidiidae, Phyllodesmium spp.:
5(2):243-258. Piseinotecus:
5(2):183-184. Pisidium annandalei, P
clarkeanum: 5(1):91-99. Placida den-
Oritica: 5(2):243-258. Plagiola inter-
rupta, P lineolata: 6(1):19-37. Platy-
dorididae, Platydoris cruenta, P
scabra: 5(2):243-258. Plethobasus
spp.: 6(1):19-37. Pleurehdera, P
haraldi: 5(2):215-241. Pleurobema
spp.: 6(1):19-37. Pleurobranchacea

304 AMER. MALAC. BULL. SUBJECT INDEX: 1983 - 1988

Pleurobranchaea, P maculata, P
meckelii: 5(2):215-241. Pleurobranch-
aeidae: 5(2):215-241, 243-258.
Pleurobranchella, P alba, P. nico-
barica: 5(2):215-241. Pleurobranchi-
dae: 5(2):215-241, 243-258. Pleuro-
branchidium, Pleurobranchillus,
Pleurobranchinae, Pleurobranchoides
gilchristi: 5(2):215-241. Pleurobranchus
spp.: 5(2):215-258. Plocamopherus
gulo: 5(2):183-184. P maculata,
Polycera spp., Polyceridae:
5(2):243-258. Polymesoda (Geloina)
erosa: 5(1):91-99. Potamilus spp.:
6(1):19-37. Pruvotfoilia pselliotes:
5(2):243-258. Ptychobranchus fascio-
lare, Ptychnobranchus subtentum:
6(1):19-37. Pupa spp.: 5(2):243-258.,
Quadrula spp.: 6(1):19-37. Retusa
truncata, Retusidae, Rictaxis albus,
Ringicula turtoni, Ringiculidae,
Risbecia pulchella, Robastra gracilis,
R. luteolineata, Rostanga muscula,
Rostangidae: 5(2):243-258. Roya, R.
spongotheras: 5(2):215-241.
Sacoglossa: 5(2):243-258. Scalptia
spp.: 2:57-61. Scaphander punc-
tostriatus, Scaphanderidae, Sclero-
doris apiculata, S. coriacea,
Scyllaeidae: 5(2):243-258. Simpsonaias
ambigua, Simpsoniconcha ambigua:
6(1):19-37. Siphonaria: 5(2):215-241.
Smaragdinella calyculata:
5(2):243-258. Solatia: 2:57-61.
Spiricella: 5(2):215-241. Stiliger or-
natus, Stiligeridae: 5(2):243-258.
Strophitus rugosus, S. undulatus:
6(1):19-37. Stylocheilus longicauda:
5(2):243-258. Susania: 5(2):215-241.
Tambja capensis, T. morosa, Tergipe-
didae, Tergipes tergipes, Tethyidae,
Thecacera pacifica, T. pennigera:
5(2):243-258. Tonicia (Lucilina) suezi-
ensis: 6(1):115-130. Toxolasma spp.:
6(1):19-37. Trapania: 5(2):243-258.
Tridachia crispata: 4(2):232. Trigona
pellucida, Trigonaphora withrowi,
Trigonostoma spp.: 2:57-61. Trito-
gonia verrucosa: 6(1):19-37. Tritonia,
T. nilsodhneri, Tritoniidae:
5(2):243-258. Tritonium viridulum:
2:57-61. Truncilla spp.: 6(1):19-37. Tur-
ridae: 3(1)98. Tylodina: 5(2):215-241.
T. alfredensis: 5(2):243-258. T. spp.,
Tylodinella, T. trinchesii, Tylodinidae,
Umbraculacea: 5(2):215-241. Um-
braculidae: 5(2):215-241, 243-258.
Umbraculum: 5(2):215-241. U. sinicum:
5(2):243-258. U. umbraculum, Um-
brella: 5(2):215-241. Uniomerus
tetralasmus: 6(1):19-37. Union
douglasiae: 5(1):91-99. Vayssieridae:
5(2):243-258. Villosa spp.: 6(1):19-37.

AMER.

Voluta cancellata, V. nassa, V.
reticulata, V. scabriculus: 2:57-61.
Volvatella laguncula: 5(2):243-258.
Williamia: 5(2):215-241
Tectonics
Distribution Effects: 2:84-85
Temperature Tolerance
Corbicula fluminea: 3(1):94. Macoma
balthica: 1:90
Teratology
Acochlidiacea: 5(2):303-306
Territoriality
Lottia gigantea: 2:80
Testicular Histology
Tarebia granifera: 1:95-96
Thin Sectioning, age determination
Lasmigona subviridis, Medionidus
conradicus, Pleurobema oviforme,
Villosa vanuxemi: 6(2):179-188

Threatened Species
Obovaria subrotunda: 3(1):105

Torsion
Garstang Theory: 1:89

Toxicology
Bivalvia, Unspecified, Catostomus
commersoni, Coleoptera: S2:69-81.
Corbicula: 3(1):106-107; S2:41-45,
47-52, 63-67, 83-88, 95-98. C.
fluminea: S2:69-81, 133-142. Crasso-
Strea virginica: S3:31-36, 41-49,
59-70. Cyprinus carpio, Diptera,
Ephemeroptera, Gambusia affinis,
Gastropoda, Unspecified, Hydro-
psyche, Ictalurus punctatus, lsonychia,
Lepomis macrochirus: S2:69-81.
Melampus bidentatus: 4(2):236-237.
Morone chrysops, Nitocris, Notropsis
spilopterus, Physa sp., Stenomena,

MALAC. BULL. SUBJECT INDEX: 1983 - 1988 305

Trichoptera: S2:69-81. Unionidae,

Unspecified: 3(1):106-107
Trapping

Octopus spp.: 6(1):45-48
Ulcers

Octopus briareus, O. joubini: 2:93-94
Vision

Octopus maya, O. vulgaris: 2:92
Visual Cues

Littorina irrorata: 2:78
X-Ray Analysis, Dispersive

Tegula sp.: 1:102
X-Ray Microanalysis

Crassostrea rhizophorae: 1:102
Zooxanthellae

Tridacna sp.: 2:83. Turbinaria

(passim): 5(2):185-196

th two additional copies for review
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aragraph. The generic name can be ab-
remainder of of the Paragraph as follows:

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Lif Skeletal Growth of Aquatic Organisms, D.
4 __C. Rhoads and R. A. Lutz, eds. pp. 23-67.
Plenum Press, New York.

Illustrations should be clearly detailed and readily
‘reproducible. Maximum page size for illustrative purposes is
17.3 cm x 21.9 cm. A two- column format. is used with a
single column being 8.5: cm wide. All line drawings should be
in black, high ‘quality ink. Photographs must be on glossy,

_ high contrast paper. All diagrams must be numbered in the

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AMERICAN
MALACOLOGICAL
BULLETIN

VOLUME 7 1989 NUMBER
‘%
CONTENTS ‘
wd ©
Campanile revisited: implications for Cerithioidean phylogeny. rag f
RICHARD S. HOUBRICK . ©. 22... ed ee ec ees ay, seat Sipe he SAP a pees lagers

Genetic consequences of partial self-fertilization on populations of
~~ Liguus fasciatus (Mollusca: Pulmonata: Bulimulidae).
Pe UM Leone ede ey pe ie he rt ee a Se ee, (EMR

Mechanical wear of radular denticle caps of Acanthopleura granulata (Gmelin,
1791) (Polyplacophora: Chitonidae). ROBERT C. BULLOCK..............................

Behavior, body patterning, growth and life history of Octopus briareus
cultured in the laboratory.
ROGER T. HANLON and MARTIN R. WOLTERDING.......................-..0.2.0020...

The ecology of Octopus briareus Robson in a Bahamian saltwater lake.
EA ATE ARONSON Gres Hotta. 2S ys kA eee ee a hao PhS SG hs whe ei and shee

An Atlantic molluscan assemblage dominated by two species of
Crassinella (Bivalvia: Crassatellidae). WILLIAM G. LYONS .......................-......

Temporal variation in microstructure of the inner shell surface of
Corbicula fluminea (Bivalvia: Heterodonta).
ANTONIETO TAN TIU and ROBERT S. PREZANT ...................0. 20.00.00. 000.0005.

The functional morphology of the organs of the mantle cavity of Batissa

violacea (Lamarck, 1797) (Bivalvia: Corbiculacea).

Pea TeM COTO Nr Ce Vwi eG wit ts. na ce haut. Rod Pao eRe PR.
Bivalves in the genus Corbicula (Bivalvia: Corbiculidae) in the

Soviet Union with a catalogue of type materials in the Zoological

Institute, Academy of Sciences of the U.S.S.R., Leningrad.

Peete GOUT IEG ue el A oh am heed bale gee oo ee tof
Mba AIC DOU ae St ce ee aN Gers ke aa nO Srey is Br ake Me aye Lee NE gn

PICT GERMCINS a Saree il cae Be he Es ac re pe lo at dV et oe Pw og

Whe NY BERT SIAM, hee 8 ys GRE OA oy Sie 5 Ba fos OR SRL HI earl SAS on ct a at Ca en i Se Se ee eg

1

AMERICAN MALACOLOGICAL BULLETIN

EDITOR-IN-CHIEF

ROBERT S. PREZANT
Department of Biology
Indiana University of Pennsylvania
Indiana, Pennsylvania 15705

MELBOURNE R. CARRIKER
College of Marine Studies

University of Delaware

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GEORGE M. DAVIS

Department of Malacology

The Academy of Natural Sciences
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American Malacologists, Inc.
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JOHN A. ALLEN
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Texas Christian University
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University of Michigan
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Gulf Coast Research Laboratory
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University of Sheffield
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+ Sorin

RONALD B. TOLL
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Los Angeles County Museum
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American Museum of Natural History
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ISSN 0740-2783

W. D. RUSSELL-HUNTER :

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THOMAS R. WALLER |
Department of Paleobiology
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University of Texas
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JOSEPH HELLER
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ALAN M. KUZIRIAN

NINCDS-NIH at the

Marine Biological Laboratory
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University of Southern Maine
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Florida State University
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ANDREW C. MILLER
Waterways Experiment Station
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BRIAN MORTON
University of Hong Kong
Hong Kong

JAMES J. MURRAY, JR.
University of Virginia
Charlottesville, Virginia, U.S.A.

RICHARD NEVES

Virginia Polytechnic Institute
and State University

Blacksburg, Virginia, U.S.A.

JAMES W. NYBAKKEN
Moss Landing Marine Laboratories
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WINSTON F. PONDER
Australian Museum
Sydney, Australia

CLYDE F. E. ROPER
U.S. National Museum
Washington, D.C., U.S.A.

NORMAN W. RUNHAM
University College of North Wales
Bangor, United Kingdom

AMELIE SCHELTEMA
Woods Hole Oceanographic Institution
Woods Hole, Massachusetts, U.S.A.

ALAN SOLEM
Field Museum of Natural History
Chicago, Illinois, U.S.A.

DAVID H. STANSBERY
Ohio State University
Columbus, Ohio, U.S.A.

FRED G. THOMPSON
University of Florida
Gainesville, Florida, U.S.A.

THOMAS E. THOMPSON
University of Bristol
Bristol, United Kingdom

NORMITSU WATABE
University of South Carolina
Columbia, South Carolina, U.S.A.

KARL M. WILBUR
Duke University
Durham, North Carolina, U.S.A.

Cover. Mantle cavity organs of Batissa violacea (Lamarck, 1797), the right valve of which is seen here in lateral view, are discussed in an

article by Morton in this volume, pages 73-80.

THE AMERICAN MALACOLOGICAL BULLETIN is the official journal publication of the American Malacological Union.

AMER. MALAC. BULL. 7(1)
April 1989

CAMPANILE REVISITED: IMPLICATIONS FOR
CERITHIOIDEAN PHYLOGENY

RICHARD S. HOUBRICK
DEPARTMENT OF INVERTEBRATE ZOOLOGY
NATIONAL MUSEUM OF NATURAL HISTORY

SMITHSONIAN INSTITUTION
WASHINGTON, D. C. 20560, U. S. A.

ABSTRACT

Aspects of the anatomy of Campanile symbolicum Iredale, the sole survivor of the large cam-
panilid lineage, are reexamined and compared with data derived from past and recent studies of this
aberrant gastropod. These data provide evidence to support a new systematic placement of Campanile
at the base of, but outside the Cerithioidean clade. The family Campanilidae, is herein raised to super-
familial rank, Campaniloidea Douville, 1904, and is regarded as an early, major radiation off the stem
that gave rise to Cerithioidea and Caenogastropoda.

The abberant, relictual taxon, Campanile symbolicum
Iredale, 1917, stands apart from all other Recent
Caenogastropoda by many unusual conchological and
anatomical features. These were first noted in an anatomical
and systematic study in which aspects of the ecology and
reproductive biology of this marine gastropod were also
represented (Houbrick, 1981). The paleontology and radiation
of the family Campanilidae Douville, 1904 were also described
and the taxonomy of the group outlined.

The Campanilidae was a large, diverse, complex group,
that attained its apogee during the early Tertiary, and is best
known from the Paris Basin Eocene fauna. The campanilid
radiation was extensive and comprised diverse genera and
numerous species. Fossils are known from the Indo-Pacific
and western Atlantic, as well as from many European Tethyan
sites, where the group was particularly diverse (Houbrick,
1981). Some taxa attained sizes of up to a meter in length,
and several Caribbean Eocene Campanile fossils of even
greater lengths, have been recently described and illustrated
(Jung, 1987).

The salient characters defining the sole living represen-
tative of the group, Campanile symbolicum, were recently sum-
marized and a cladogram illustrating its position relative to
other cerithioidea taxa was presented (Houbrick, 1988:115, fig.
2). Since that work, new studies on prosobranch phylogeny,
that include other aspects of Campanile anatomy (Haszprunar,
1985; 1988; Salvini-Plawen and Haszprunar, 1987; and
Houbrick, 1988), and comprehensive ultrastructural studies
of its spermatozoa (Healy, 1983; 1986a, b), have been pub-
lished. These studies examined specific anatomical traits in

more detail, and have provided new characters and additional
data enhancing our understanding of the systematic place-
ment of this strange gastropod.

An opportunity to restudy living Campanile specimens
in Albany, Western Australia, allowed me to check my previous
work (Houbrick, 1981) for accuracy as well as to make several
new observations. The original study (Houbrick, 1981) was
conducted during the Australian winter, while the present one
was made in the summer of 1988 (Jan). The new study and
data from recently published findings of the above mentioned
authors, reinforce the concept that Campanile is indeed an
aberrant gastropod, difficult to place within the framework of
conventional molluscan systematics. Reevaluation of the
original findings, additional data compiled from the recent
literature, and new anatomical information obtained from the
present investigation, all indicate that the phylogenetic rela-
tionship of the family Campanilidae to the superfamily
Cerithioidea is in need of critical examination and reassess-
ment, and have prompted this paper.

MATERIALS AND METHODS

Live specimens of Campanile symbolicum lredale, 1917,
were collected in Princess Royal Harbour, Albany, Western
Australia, in shallow water amongst rocks and sand associated
with Possidonia grass beds, during January, 1988. Shells were
cracked with a vice, and the animals extracted and relaxed
in a 7.5% MgClz solution isotonic with seawater prior to dissec-
tion. Radulae and tissues prepared for critical point drying
were examined with an Hitachi S-570 Scanning Electron

American Malacological Bulletin, Vol. 7(1) (1989):1-6

;

2 AMER. MALAC

Microscope (Figs. 1-6). The egg mass was photographed
under a Wild M-5 dissecting microscope (Figs. 7-8). Material
for histological examination was fixed in Bouin’s fixative,
embedded in paraffin, and sectioned at 5 um. Sections were
stained in Mallory’s Triple Stain (Figs. 9-10). The characters
derived from this study were compared with my cladogram

Figs. 1-6. Scanning electron micrographs of Campanile symbolicum anatomy (all USNM 867015, Princess Royal Harbour, Albany, West Australia).
Fig. 1. Radula with right marginal teeth spread back. Fig. 2. Detailed view of rachidian and lateral teeth. Fig. 3. Spirally arranged leaflets
removed from anterior liver duct. Fig. 4. Detail of ribs on liver duct leaflet showing ciliated epithelium. Fig. 5. Detail of surface of pad-like

. BULL. 7(1) (1989)

on cerithioidean phylogeny (see Houbrick, 1988, Fig. 2) for
congruence and used as an independent test of my original
conclusions about the placement of Campanile. Voucher
specimens from Albany, Western Australia (USNM 867015),
have been deposited in the National Museum of Natural
History, Smithsonian Institution.

fold emerging from spiral caecum, showing densely packed papillae. Fig. 6. Ciliated folds of anterior hypobranchial gland.

HOUBRICK: CAMPANILE 3

RESULTS

The basic anatomy of Campanile symbolicum has been
set forth in a previous paper (Houbrick, 1981) and these
original findings verified by dissections made during this
study. New comments clarifying past descriptions, several cor-
rections, and new anatomical observations follow.

The arrangement of the dark brown digestive gland and
light colored gonad in the visceral whorls of Campanile is
unusual. The wide gonad (? ovotestis) is sharply demarcated
from the digestive gland on the peripheral surface of the ex-
ternal anterior visceral whorls, exclusive of the kidney and
stomach, presenting a banded appearance not normally seen
in other prosobranchs. In addition, the gonad appears to be
dispersed throughout the interior of the digestive gland.

One of the more interesting and unusual features of
Campanile is the alimentary system, which has features that
are unique among Caenogastropoda. Deep epithelial folds
line the lips, and the thick, four-layered ultrastructure of the
large, stout jaws (see Houbrick, 1981:274, fig. 2d,e) does not
occur among other cerithioideans, or as far as is known,
among other prosobranchs. As pointed out previously
(Houbrick, 1981:274-275), the radular ribbon (Fig. 1) is very
wide and robust, but unusually short for so large an animal,
attaining a length only about 8% of the shell length. The rachi-
dian tooth is notable in having a very large, broad, central cusp
with only weak traces of minor denticles (Fig. 2). The lateral
teeth are similar but have a small inner denticle (Fig. 2). The
interior buccal cavity is lined with deeply folded, glandular
tissue, which greatly increases its surface area. The large,
so-called esophageal pouches are unusual structures and it
is doubtful that they are homologous with the esophageal
pouches described by Fretter and Graham (1962:26) in Lit-
torina; hence, my hesitation in using the same name for these
structures. In Campanile, the pouches differ from those of Lit-
torina in being more internally complex and highly muscular,
in having a short, narrow, highly constricted connection to the
buccal cavity, and in being lined with thick, dark-staining glan-
dular tissue of unknown function (Houbrick, 1981:275, fig. 7f).
The salivary glands are tiny relative to the size of the snail,
and are located well anterior to the nerve ring. The existence
of a muscular, transverse septum behind the nerve ring, com-
pletely dividing the cephalic haemocoel between the anterior
esophagus and the mid-esophagus, is confirmed. As noted
previously (Houbrick, 1981:275), a transverse septum is known
only in trochaceans (archaeogastropods), but not in
caenogastropods. The mid-esophagus of Campanile is highly
unusual in that it is surrounded by a thick layer of very loose
connective tissue which is in turn surrounded by a thin layer
of muscular tissue (see Houbrick, 1981:275, fig. 7e).

The morphology of the large stomach of Campanile,
one of its strangest features, sets it apart from those of all
other caenogastropods. The drawing of the stomach
presented by Houbrick (1981:fig. 5b) is essentially accurate,
’ but a few points need clarification: 1) The fold emerging from
the spiral caecum (ff on drawing) is a large pad-like structure
comprised of densely packed papillae having a ciliated sur-
face (Fig. 5); 2) The so-called gastric shield in the drawing

(gs) is not a gastric shield, but a raised, ciliated pad adjacent
to the large fold (ff) and to the sorting area; 3) The grooved
sorting area in the drawing (gsa) is cuticularized, and is pro-
bably homologous to the cuticularized part of the anterior
chamber and perhaps the gastric shield of other caeno-
gastropods; 4) There are two openings (ducts) into the anterior
lobe of the digestive gland rather than one, and contrary to
the original description (Houbrick, 1981:276), the so-called
“pit” with spirally arranged leaflets is not blind, but branches
deeply into the digestive gland. Both openings are large, and
the unusual spirally arranged leaflets comprising the anterior
digestive gland duct branch deeply into the far anterior lobe
of the digestive gland. The illustration of these leaflets in the
original description (Houbrick, 1981:276, fig. 5, sl) was inade-
quate, and is here augmented by scanning electron micro-
graphs (Figs. 3-4). The leaflets are largest at the conical
shaped opening to the digestive gland and become pro-
gressively smaller as they spiral downward. Each of the
leaflets is transversely ribbed, presenting a veined appearance
and is entirely covered with small cilia (Fig. 4); 5) Although
a shallow, rudimentary style sac is present, there is a raised
cuticular area posterior to the style sac instead of a conven-
tional gastric shield and the crystalline style is absent. The
stomachs of freshly taken snails showed no trace of a
crystalline style: only a short protostyle (sensu Morton,
1967:112) was present.

The digestive gland is very large, comprising 4-5
whorls, and is a very dark color due to the great numbers of
deep brown concretions (Fig. 10, dg), which are found in the
basal parts of the cells and which loosen and fall out when
the gland is cut, and darkly stain preserving fluids.

The large, saddle-shaped, dark tan kidney is a con-
spicuous external feature of the animal. There are two main
parts to the kidney: an elongate, solid section, comprised of
many fine lamellae (Fig. 9, k), covers the pericardium dorsal-
ly and posteriorly and extends posteriorly to overlie the
posterior gonoduct and posterior mantle cavity; it has a
spacious anterior cavity or kidney sac (Fig. 9, ks), adjacent
to the kidney opening (Fig. 9, ko); the other part of the kidney,
of looser, spongy consistency due to larger lamellae and
numerous lumina, is seen in section to be filled with tiny ex-
cretory granules; it is adjacent to and surrounds the anterior
stomach. This was not identified as a separate part of the
kidney in the original description (Houbrick, 1981:276). A third,
lighter pigmented part of the kidney, thought to be the
nephridial gland (Houbrick, 1981:276), borders the pericar-
dial sac, but histologically does not appear to consist of dif-
ferent tissue than that of the main kidney.

DISCUSSION

Although the large, well-developed, bipectinate
osphradium of Campanile is similar to those of rachiglossate,
predatory neogastropods, herbivory has been reconfirmed.
Stomach contents and a large food bolus taken from the
anterior liver duct consisted of large pieces of Possidonia
seagrass as well as coarse fragments of foliated and ar-
ticulated algae, which were primarily Cladophora, but also

4 AMER. MALAC. BULL. 7(1) (1989)

a
4
a

aed

g ay?

a

2

filamentous threads between egg chambers (bar = 0.6 mm). Fig. 9. Section through kidney sac showing kidney tissue (k), kidney sac (ks),and
kidney opening (ko) to mantle cavity (mc); (bar = 0.25 mm). Fig. 10. Section through digestive gland (dg) and testis (t) showing connective
tissue (ct) and a duct of the digestive gland (dgd). Dark round objects are digestive gland concretions (bar = 1 mm).

some Specelaria. Algal pieces are probably manipulated and
compressed by the large pad-like structures in the stomach
to form a bolus prior to its movement into the liver ducts. The
unusual, large, branched, structure of the interior liver ducts
(Fig. 10, dgd) suggests that the bolus of algal fragments is
pushed deeply into them. The many unusual features of the
alimentary system indicate that the feeding ecology and
digestive physiology of Campanile would be an interesting
study.

In the discussion of my original paper (Houbrick,
1981:283-289) | proposed and justified familial status for Cam-
panile, and allocated Campanilidae to the superfamily
Cerithioidea; however, | noted that the many unique and
unusual anatomical features of the only living representative
of the family, Campanile symbolicum, do not conform to the
normal cerithioidean groundplan (see Houbrick, 1988 for
detailed description of Cerithioidea), and that some characters
suggest affinities with neogastropods (rachiglossa) and

HOUBRICK: CAMPANILE 5

opisthobranchs.

In a recent study of cerithioidean phylogeny (Houbrick,
1988) in which 58 characters comprising 134 character states
were used to generate cladograms of 15 cerithioidean families,
the Campanilidae consistently fell out at the bases of the
various trees generated, irrespective of outgroups used or of
interpretations of multistate character polarity. Among all
cerithioidean families, Campanilidae was consistently the
most primitive taxon and was at the base of the final, most
parsimonious cladogram (Houbrick, 1988: fig. 2). Eleven
autapomorphies defining the Campanilidae were identified,
and it was remarked that Campanile would suffice as a good
outgroup for all other cerithioidean taxa, and that it occupied
an isolated position at the base of the Cerithioidea clade
(Houbrick, 1988).

The new characters set forth in this paper reinforce the
basal position of Campanile on the original cladogram
(Houbrick, 1988: fig. 2). Comprehensive studies of the eusper-
matozoa and paraspermatozoa of Campanile symbolicum by
Healy (1983, 1986a, 1986b) indicate that major, significant dif-
ferences exist between spermatozoa of the Campanilidae and
other cerithioidean taxa, confirming its unique status among
prosobranchs. The nucleus of the euspermatozoon of Cam-
panile is three times the length of euspermatozoan nuclei of
all other investigated cerithioideans. Although the basic struc-
ture of Campanile euspermatozoa resembles that of many
other mesogastropods, the midpiece region exhibits unusual
and possibly unique features (see Healy, 1986b:213). In ad-
dition, Campanile differs from all other cerithioidean taxa
studied in having two types of paraspermatozoa, both with
a head (acrosome-like structure) and 2-3 tails. These two types
of paraspermatozoa are nucleate, and non-nucleate (Healy,
1986b:207-209). Despite these significant differences, Healy
(1986b:214-216) pointed out that Campanile paraspermatozoa
also share a number of important features with those of the
Cerithiidae, Potamididae, Turritellidae, and Planaxidae, and
that in many respects the anatomy and sperm morphology
of Campanile bridge the gap between the Cerithioidea and
the remainder of the Caenogastropoda. He concluded that
sperm morphology indicates that the Campanilidae occupy
an isolated position within the Cerithioidea and that they pro-
bably diverged at an early stage from the primitive cerithioi-
dean stock in which sperm dimorphism was established.
Healy’s (1986b) position was adopted by Ponder and Waren
(1988), who considered Campanile to be an aberrant
cerithioidean.

Haszprunar (1985:24) called attention to the fact that
within the Prosobranchia, only Valvata and Campanile have
chalazae. Salvini-Plawen and Haszprunar (1987:762, fig. 4)
allocated the Campanilidae to the Caenogastropoda, but as
incertae sedis, and suggested that the group is a ‘‘subsequent
offshoot of intermediate grade’ between the Caenogastro-
poda and Allogastropoda (sensu Haszprunar, 1984). Thus,
they considered Campanilidae to be distinct from and not
directly ancestral to Cerithioidea, and suggested that Cam-
panilidae is an intermediate group of caenogastropods shar-
ing some characters with euthyneurans, which they called
“‘Pentaganglionata’’. Salvini-Plawen and Haszprunar

(1987:765) placed emphasis on the presence of chalazae and
on the anterior folds (presumed respiratory lamellae) of the
hypobranchial gland epithelium as evidence for the transi-
tion between the prosobranch and heterobranch grades. The
following caveats about this evidence should be noted: 1) In
my original paper | mentioned that the string-like connections
in Campanile spawn masses (see Figs. 7-8), are merely be-
tween the mucous capsules forming egg chambers (each con-
taining 1-3 eggs) and not between individual eggs. | pointed
out that these connections may not be homologous with the
true chalazae of opisthobranch spawn (Houbrick, 1981:285),
which join individual eggs; 2) reexamination of the anterior
hypobranchial gland casts doubt on its role as a primitive
respiratory lamellae. The folds appear to be more poorly de-
fined and less prominent (see Fig. 6) than in my original
description (Houbrick, 1981:274, fig. 4, A, /hg), and it is doubtful
that their function is respiratory.

Haszprunar (1988) recently emphasized that Campanile
has certain characters of the allogastropod-euthyneuran line;
i.e., a genital system with isolated receptaculum, and a
gelatinous egg mass with chalazae-connected eggs. In the
same paper, he also cited his fine-structural studies of the
Campanile osphradium (without giving details), which he
stated demonstrate a major difference from osphradia of other
caenogastropod groups, and which indicate affinities with the
Architectonicidae and primitive Euthyneura. Haszprunar
(1988) thus concluded that Campanile probably represents a
first step towards the euthyneurous level of organization.
However, Ponder and Waren (1988) pointed out that un-
doubted fossil opisthobranchs are known from as far back
as the Carboniferous, suggesting that euthyneurans arose
long before any Campanile-like gastropods.

The new observations of Campanile anatomy and
reanalysis of anatomical characters, plus work on sperm mor-
phology (Healy, 1983; 1986a; 1986b) and on other aspects
of Campanile anatomy (Haszprunar, 1985; 1988; Salvini-
Plawen and Haszprunar, 1987), all confirm the unusual posi-
tion of Campanilidae relative to Cerithioidea, and Caeno-
gastropoda-Allogastropoda, and perhaps to Euthyneura (Pen-
taganglionata), and provide evidence arguing for a major
reevaluation of its systematic placement.

The Campanilidae should no longer be considered as
cerithioidean gastropods. Campanile has many notable and
significant non-cerithioidean features including: a calcified,
pitted periostracum, a complex jaw ultrastructure; buccal
pouches; digestive gland openings with spirally arranged
leaflets; a transverse septum dividing the cephalic hemocoel
from the midesophagus; an unusual arrangement of muscle
and connective tissue surrounding the midesophagus; a
lamellate albumen gland; a seminal receptacle in both sexes;
an isolated, posterior seminal receptacle positioned in the
pericardial sac; possible protandry; two kinds of para-
spermatozoa; a possibly unique form of the euspermatozoan
midpiece; egg chambers linked by chalazae-like strings; lack
of hyaline capsules around the eggs; and a short, oval, bipec-
tinate osphradium with unique epithelium. These characters,
all of which are autapomorphic among cerithioideans, plus
other minor anatomical features, are of sufficient importance

6 AMER. MALAC. BULL. 7(1) (1989)

and weight to justify removal of Campanile, family Cam-
panilidae, from Cerithioidea, and to raise it to superfamilial
status (Campaniloidea Douville, 1904). This is contrary to my
original classification (Houbrick, 1981:286). A similar view was
arrived at independently by Ponder and Waren (1988), who
stated that Campanile was an aberrant cerithioidean and had
little to do with heterobranch evolution.

Salvini-Plawen and Haszprunar (1987) and Haszprunar
(1988, in press) have suggested an intermediate, outlying posi-
tion for Campanilidae between the rest of the Caeno-
gastropoda and the Allogastropoda and Euthyneura, based
on three anatomical characters. Two of the so-called
euthyneuran features cited for Campanile by these authors,
chalazae and respiratory folds of the hypobranchial gland,
are based on equivocal evidence and are somewhat
speculative (see above). The third and possibly best of these
characters, the unique osphradial epithelium, is stated to have
features in common with Architectonicidae and primitive
euthyneurans, but Haszprunar (1988, in press) himself has
noted that the osphradium also has several peculiarities of
its own, and is unique among Gastropoda. As he does not
give details, it is impossible to appraise and judge these
osphradial characters. In my opinion, too much significance
has been given to the putative euthyneuran characters in
Haszprunar’s cladogram and resulting classification (1988).
Until the ambiguities about the above characters are resolved,
less significance and emphasis should be accorded to Cam-
panile as a ‘‘connecting link’’ between Caenogastropoda and
Euthyneura.

| agree with Haszprunar’s (1988) suggestion that the
Loxonomatoidea-Cerithioidea stem-group probably gave rise
to the Caenogastropoda, and concur that Campanilidae be
given superfamily status. However, | believe it best to regard
Campaniloidea as an early, major radiation from the
mainstream of the stem-group that gave rise to modern
Cerithioidea and Caenogastropoda.

ACKNOWLEDGMENTS

Much of this research was done at the invertebrate workshop
held at Albany, Western Australia, 1988, and sponsored by the Western
Australian Museum, Perth. | am grateful to Dr. F. Wells for logistic
support and for assistance with shipping specimens collected in the
field, | thank Dr. W. F. Ponder, Australian Museum, Sydney, for “‘look-
ing over my shoulder”’ during dissections and sharing ideas and com-
ments with me about aspects of Campanile anatomy. Support for this

project and the field work was provided by a Smithsonian Research
Opportunities Fund. | thank the staff of the Smithsonian’s Scanning
Electron Microscope Laboratory for their assistance, and Mr. V. Krantz,
Smithsonian Photographic Services, for developing negatives and
making prints. Riidiger Bieler, Delaware Museum of Natural History,
and Robert Hershler, National Museum of Natural History, Smithson-
ian Institution, kindly read drafts of this manuscript and offered
valuable comments.

LITERATURE CITED

Fretter, V. A. and A. Graham. 1962. British Prosobranch Molluscs, their
Functional Anatomy and Ecology. Ray Society, London. 755 pp.

Haszprunar, G. 1985. The Heterobranchia - a New Concept of the
Phylogeny of the Higher Gastropoda. Zeitschrift fur zoologische
Systematik und Evolutionsforschung, 23(1):15-37.

Haszprunar, G. 1988. On the Origin and Evolution of Major Gastropod
Groups, with Special Reference to the Streptoneura (Mollusca).
Malacological Review, 1988, Supplement 4 (in press).

Healy, John M. 1983. Ultrastructure of Euspermatozoa of Cerithia-
cean Gastropods (Prosobranchia: Mesogastropoda). Journal
of Morphology 178:57-75.

Healy, John M. 1986a. Ultrastructure of Paraspermatozoa of Cerithia-
cean Gastropods (Prosobranchia: Mesogastropoda).
Helgolander Meeresuntersuchungen, 40:177-199.

Healy, John M. 1986b. Euspermatozoa and Paraspermatozoa of the
Relict Cerithiacean Gastropod, Campanile symbolicum (Pro-
sobranchia, Mesogastropoda). Helgofander Meeresunter-
suchungen, 40:201-218.

Houbrick, Richard S. 1981. Anatomy, Biology and Systematics of Cam-
panile symbolicum with Reference to Adaptative Radiation of
the Cerithiacea (Gastropoda: Prosobranchia). Malacologia,
21(2-3):263-289.

Houbrick, R. S. 1988. Cerithioidean Phylogeny. Malacological Review,
1988, Supplement 4 (in press).

Jung, P. 1987. Giant Gastropods of the Genus Campanile from the
Caribbean Eocene. Ecologae Geologicae Helvetiae,
80(3):889-896.

Morton, J. E. 1967. Mollusca. Hutchinson University Library. London.
244 pp.

Ponder, W. F. and A. Waren. 1988. Classification of the
Caenogastropods and Heterostropha - a List of the Family-
Group Names and Higher Taxa. Malacological Review, 1988,
Supplement 4:288-326.

Salvini-Plawen, L. v. and G. Haszprunar. 1987. The Vetigastropoda
and the Systematics of Streptoneurous Gastropods (Mollusca).
Journal of Zoology, London, 211:747-770.

Date of manuscript acceptance: 25 August 1988

GENETIC CONSEQUENCES OF PARTIAL SELF-FERTILIZATION

ON POPULATIONS OF LIGUUS FASCIATUS
(MOLLUSCA: PULMONATA: BULIMULIDAE)

DAVID M. HILLIS
DEPARTMENT OF ZOOLOGY
THE UNIVERSITY OF TEXAS
AUSTIN, TEXAS 78712, U.S.A.

ABSTRACT

Reproductive modes of the highly polymorphic Florida tree snail, Liguus fasciatus (Muller), were
investigated by laboratory breeding experiments and field study. Variation of glucose-phosphate
isomerase and shell phenotypes was assessed. The laboratory crossings demonstrated that partial
self-fertilization does occur in this species, but too few informative crosses were performed to estimate
the frequency of self-fertilization. A transect study through two populations that have recently come
into contact demonstrated high population substructure (Fst = 0.437) across short distances. Levels
of heterozygosity in subpopulations along 20 m sections of the transect were used to estimate levels
of self-fertilization. Estimates ranged from 46% to 94% self-fertilization, with a mean of 69%. The
genotypic frequencies of subpopulations did not differ significantly from expected frequencies assuming
the mean estimate of 69% self-fertilization, but did differ significantly from expected frequencies assuming
Hardy-Weinberg equilibrium with no self-fertilization. Partial self-fertilization appears to be largely respon-
sible for the low within-population variation compared to the high among-population variation of this

species.

Tree snails of the genus Liguus are noted for their mor-
phological diversity among populations (Clench, 1946, 1954,
1965; Pilsbry, 1946). In Florida, approximately 58 named
varieties of Liguus fasciatus (Muller) occur (Roth and Bogan,
1984), many of which are restricted to single tropical hard-
wood hammocks in the Everglades and Florida Keys (Deisler,
1982). In spite of this high morphological diversity in L.
fasciatus, allozymic variation is very low among and within
populations of this species (Hillis et a/., 1987). Furthermore,
populations of L. fasciatus deviate significantly from Hardy-
Weinberg expectations at variable loci because of marked
heterozygote deficiencies (Hillis et a/., 1987).

Although geographic patterns of phenotypic shell varia-
tion have been studied extensively in Liguus fasciatus (Pilsbry,
1899, 1912, 1946; Deisler, 1982; Roth and Bogan, 1984), very
little is Known about the inheritance of these traits or reproduc-
tion in this species. Roth and Bogan (1984) proposed a system
for describing morphological variation in L. fasciatus that con-
sisted of twelve characters, each with two to four states. They
stated that they chose characters ‘‘...in which the alternate
states can be seen to segregate in randomly selected
material.’ However, Hillis et a/. (1987) suggested that these
characters are not independent, and that many fewer than

twelve loci are probably responsible for the observed
phenotypic variation of shells. Furthermore, although most
past authors (e.g. Brown, 1978; Young, 1960) have assumed
that L. fasciatus is an obligate outcrosser, Hillis et a/. (1987)
suggested that partial self-fertilization might account for some
of the patterns of genetic variation seen among populations
of this hermaphroditic species. Self-fertilization and outcross-
ing are both common modes of reproduction in gastropods,
and a few species contain some populations that are self-
fertilizing and others that are outcrossing (McKracken and
Selander, 1980). Other species are facultatively self-fertilizing
and self-fertilize when mates are unavailable, and in at least
one species reproduction following copulation is either by self-
fertilization or outcrossing (McKracken and Selander, 1980).
However, partial self-fertilization (a single clutch containing
both self-fertilized and outcrossed eggs), as suggested for L.
fasciatus (Hillis et a/., 1987), has not been demonstrated
among gastropods.

This study was undertaken to determine the mode of
reproduction and its consequences on genetic variation in
Liguus fasciatus. Laboratory and field studies were designed
to determine if partial self-fertilization occurs, and if so, at what
frequency. In addition, a population was examined to

American Malacological Bulletin, No. 7(1) (1989):7-12

7

8 AMER. MALAC. BULL. 7(1) (1989)

determine the extent of genetic substructure as well as the
effects of possible self-fertilization on heterozygosity, allozymic
variation, and phenotypic variation of shells.

MATERIALS AND METHODS

ELECTROPHORETIC METHODS

Standard procedures of horizontal starch gel elec-
trophoresis were followed (Selander et a/., 1971; Hillis, 1985).
Digestive glands of Liguus fasciatus were ground and diluted
1:1 in 0.01 M tris-0.001 M EDTA-.001 M 2-mercaptoethanol, pH
7.5. Homogenates were centrifuged at 7,000 g for 5 min, after
which the supernatants were refrozen at -85°C. A buffer
system of 175 mM tris-17.5 mM boric acid-2.75 mM EDTA, pH
9.1 was used. Gels were prepared from 50% Sigma starch
(lot 85F-0010) and 50% Otto Hiller electrostarch (lot 392). Gels
were electrophoresed for 12 hr at 12.5 Vicm. Histochemical
staining for glucose-phosphate isomerase (E.C. 5.3.1.9; GP)
followed Harris and Hopkinson (1976). This enzyme was the
only variable locus of the 24 allozyme loci surveyed in L.
fasciatus by Hillis et a/. (1987).

BREEDING STUDY

Between 18 January and 5 July 1986, 60 specimens
of Liguus fasciatus were collected from hammocks in the
Pinecrest region, Big Cypress National Preserve, and near
Long Pine Key, Everglades National Park, Florida, for cap-
tive breeding experiments. Mating in this species begins in
late July or early August in these regions (Jones, 1954). Pairs
of L. fasciatus remain together for several days after mating,
so the beginning of the breeding season can be easily ascer-
tained. In summer 1986, the study populations were observed
at least twice weekly, and the first mated pairs were found
during the first week of August. Therefore, all of the specimens
used in the captive breeding study were collected at least one
month prior to the breeding season. Specimens from single
populations were paired at random and kept in isolation in
plastic boxes (10 cm x 20 cm x 30 cm) with 3-4 cm of decayed
leaves and hammock soil. Snails were fed with lichen-covered
branches supplemented with a mixture of cornstarch, oatmeal,
spinach, vitamins, and calcium carbonate. Snails were main-
tained at approximately 25°C, and were sprayed with water
5 times per week until eggs were deposited (24 Sept - 5 Oct).
During egg deposition, the egg-producing individuals were
marked. After eggs had been deposited, cages were sprayed
with water at approximately two week intervals until hatching
occurred (Jan - Feb 1987). Parental snails and offspring were
then examined for variation at the glucose-phosphate
isomerase locus as described above.

FIELD STUDY

The study site was located near Pinecrest, Big Cypress
National Preserve, Monroe County, Florida. Pinecrest ham-
mocks (PC) 16 and 16a (numbering system follows Pilsbry,
1946) were separated by a narrow channel of water until the
1960’s or 1970’s (Hillis et a/., 1987; Fig. 1). Prior to connec-
tion of these hammocks, Liguus fasciatus in PC 16 were of

PC 16 PC |6a

ro)
fo)

504

Percent of sample

108)
°

Percent of sample

(@)

120. ‘160

Meters along transect

Fig. 1. A. Map of Pinecrest hammocks 16 and 16a, showing loca-
tion of transect. The shading around the hammocks represents the
approximate extent of recent woody growth that is seasonally flood-
ed. This growth provides a connection between the hammocks for
movement of Liguus fasciatus. B. Shell phenotypes of L. fasciatus
collected in corresponding sections of the transect shown in A. The
darkly shaded portion of the histogram represents the percentage
of the barbouri phenotype, the lightly shaded portion the aurantius
phenotype, and the white portion the walkeri phenotype. C. GPI allelic
frequencies of L. fasciatus collected in corresponding sections of the
transect shown in A. The darkly shaded portion of the histogram
represents the percentage of the F allele in the sample, and the white
portion the percentage of the S allele.

the walkeri phenotype (banded shells with pink tips), whereas
L. fasciatus in PC 16a were of the barbouri phenotype (dark
snails with white tips); a third phenotype, aurantius (orange
snails), was uncommon in both hammocks (Hillis et a/., 1987).
Fire prevention in the Pinecrest area over the past several
decades has resulted in increased woody growth around
many hammocks, and by the 1960’s or early 1970’s tree
growth (primary willows) had joined the two hammocks suffi-
ciently for movement of Liguus between PC 16 and PC 16a
(Fig. 1A). Because the two populations are also strongly dif-
ferentiated at the glucose-phosphate isomerase locus, this
site provided an opportunity to study the effects of self-
fertilization on the interaction of differentiated populations of
L. fasciastus.

A transect was constructed perpendicular to the axis
of the contact through the two hammocks (Fig. 1). Fourteen

HILLIS: PARTIAL SELF-FERTILIZATION OF TREE SNAILS 9

20 m intervals were marked along the transect, and snails
were collected from sections perpendicular to these 20 m in-
tervals. Between 20 and 44 snails were collected from each
section. For each snail, section number and morphological
phenotype were recorded; snails were then transferred to the
laboratory where each was assessed for genotype at the GPI
locus.

ANALYSIS

F-statistics were calculated using the formulae of Weir
and Cockerham (1984), which do not make assumptions con-
cerning numbers of populations, sample sizes, or
heterozygote frequencies. Indirect estimates of self-fertilization
were calculated using the method described by Hedrick
(1983). Statistical tests for goodness-of-fit and correlation were
conducted as described by Sokal and Rohlf (1981).

RESULTS

Of the 30 pairs of Liguus used in the captive breeding
experiments, 11 pairs produced clutches of eggs. In one of
these pairs, both individuals produced clutches. However, it
was determined after the pairings had been made that many
pairs came from populations that were fixed for one or the
other of the GPI alleles, so only one of the crosses was infor-
mative about self-fertilization (Table 1). In this cross, a snail
heterozygous for the two GPI alleles (FS) was mated with a
snail homozygous for the fast GPI allele (FF). The FS in-
dividual produced eggs, and the offspring expressed FF, FS,
and SS genotypes (Table 1).

In the field study, both GPI allelic frequencies and shell
phenotypic frequencies changed markedly along the transect
between PC 16 and PC 16a. The F allele of GPI increased
and the S allele decreased along this transect (from PC 16
to PC 16a), and there was a corresponding shift in frequen-
cies from mostly walkeri phenotype to mostly barbouri
phenotype (Fig. 1 and Table 2). Frequencies of heterozygotes
at GPI were considerably below Hardy-Weinberg expectations
(Table 3).

DISCUSSION

Both population substructuring and self-fertilization ap-
pear to have major effects on reduction of heterozygosity in
populations of Liguus fasciatus. Even though the transect was
divided into subpopulations just 20 m wide, variation among
subpopulations is very high (Fst = 0.479). This value is even

Table 1. GPI genotypes of offspring resulting from 12 crosses of Liguus
fasciatus.

Number Maternal Paternal Offspring

clutches genotype genotype FF FS Ss
5 FF FF 76 0 0
6 Ss SS 0 0 106
1 FS FF 7. 6 1

Table 2. Observed GPI genotypes of Liguus fasciatus from sections
along a transect through Pinecrest hammocks 16 and 16a, and
estimates of frequency of self-fertilization (S) in each section.

GPI genotype

Section Ss SF FF S
1-20 m 24 9 11 71
21-40 m 29 8 5 58
41-60 m 26 8 3 46
61-80 m AZ. 4 10 84
81-100 m 25 4 4 aT As)
101-120 m 20 vA 6 65
121-140 m 10 11 12 50
141-160 m 5 3 18 82
161-180 m 5 1 20 94
181-200 m 0 0 20 —
201-220 m 1 2 17 62
221-240 m 0 0 20 _—
241-260 m 0 0 20 —
261-280 m 0 ) 20 _—

higher than the average fixation index for self-fertilizing plants
(Fst = 0.437; Hamrick, 1983). In addition, the inbreeding
coefficient is also very high (Fis = 0.478), indicating substan-
tial self-fertilization. The reduction in individual heterozygosity
in the study population due to both of these factors (substruc-
turing and inbreeding) is quite substantial (Fj7 = 0.728).

Except for potential sperm-storage from the previous
breeding season, a possibility unsupported by data, the cap-
tive breeding data demonstrate that self-fertilization does oc-
cur in Liguus fasciatus, because only through self-fertilization
could the SS offspring result from a mating of FS x FF in-
dividuals (Table 1). However, a direct estimation of self-
fertilization frequency is not possible from the captive breeding
data because of the paucity of appropriate crosses. On the
other hand, it is possible to estimate self-fertilization frequency
from the transect study.

The proportion of progeny produced by self-fertilization
(S) can be estimated from the proportion of heterozygous in-
dividuals (H) in each of the suppopulations in the transect by
solving the equation

4pq (1-S)
2-S

where p and q are the allelic frequencies (Hedrick, 1983). This
requires the assumption that the subpopulation divisions are
small enough to account for population substructuring. Given
that individual seasonal movements of Liguus fasciatus are
typically greater than the 20 m widths of the transect sections
(Brown, 1978), this assumption is probably valid. The above
calculations were made for each of the subpopulations in
which allozymic variation was observed (Table 2). These
estimates range from 46% to 94% self-fertilization (S = 0.69,

H=

' SD = 0.154). If population substructuring is not fully ac-

counted for by the 20m transect divisions, then these
estimates of self-fertilization are somewhat inflated. An alter-
native to self-fertilization that could explain the deficiency of
heterozygotes is assortative mating. However, in polymorphic
populations of L. fasciatus mating appears to be random with

10 AMER. MALAC. BULL. 7(1) (1989)

Table 3. Expected genotypic frequencies of no self-fertilizing and 69% self-fertilizing models for sec-
tions of the PC 16-16a transect, and probabilities of the observed data fitting the expected frequen-
cies. ‘‘n.s.’ designates expected frequencies that do not differ significantly (p > 0.05) from the observed

values.
No self-fertilization 69% self-fertilization
Section SS SF FF p Ss SF FF p
1 18.5 20.1 55 <.001 23.9 9.5 10.7 ns.
2 26.0 14.1 19 <.01 29.6 68 56 n.s.
3 24.3 11.3 13 n.s. 273 5.3 43 n.s.
4 1127 14.7 46 <.001 15.5 69 8.6 ns.
5 22.1 98 1.1 <.005 246 4.7 3.7 ns.
6 16.7 13.5 2.7 <.01 20.3 6.4 63 ns.
7 7.2 16.4 9.3 <.05 11.6 78 13.6 n.s.
8 1.6 98 14.6 <.01 4.2 46 17.2 n.s.
9 1.2 8.7 16.1 <.001 3.4 4.1 18.4 <.05
10 0 0 20.0 ns. 0 0 20.0 n.s.
11 0.2 3.7 16.1 <.01 1.2 18 17.0 ns.
12 0 0 20.0 n.s. 0 0 20.0 n.s.
13 0 0 20.0 ns. 0 0 20.0 n.s.
14 0 0 20.0 ns. 0 0 20.0 ns.

respect to shell phenotype (Brown, 1978).

The expected genotypic frequencies under assump-
tions of no self-fertilization and 69% self-fertilization (the mean
of estimates from all subpopulations) are shown in Table 3
and graphically in figure 2. The observed frequencies were
tested against the expected frequencies under these two
models using a G-test (Sokal and Rohlf, 1981). All but one
of the genetically variable subpopulations differ significantly
from the expected genotypic frequencies under the assump-
tion of no self-fertilization, whereas only one of the subpopula-
tions differ significantly from the expected genotypic frequen-
cies under the assumption of 69% self-fertilization (Table 3).
The single subpopulation that differed from 69% self-
fertilization expectations differed in having even fewer
heterozygous individuals than expected. Given the number
of comparisons (10 variable subpopulations), a single depar-
ture from expectations at p = 0.05 would be expected by
chance 50% of the time, even if the model is correct.
Therefore, the transect data are in close agreement with a
self-fertilization frequency of approximately 69%.

The frequencies of GPI alleles are strongly correlated
with shell phenotypes (Fig. 3), an observation that is probably
a result of historical restriction of the barbouri phenotype and
the F allele to PC 16a, and the waj/keri phenotype and S allele
to PC 16. These two correlations are nearly equally strong
and significant: barbouri-F allele, r = 0.95, p < .001; walkeri-
S allele, r = 0.94, p < .001. The third (uncommon) phenotype,
aurantius, is not significantly correlated with either GPI allele,
which is consistent with the distribution of this phenotype in
both PC 16 and PC 16a before the contact of the two ham-
mocks. However, the distribution of the two primary
phenotypes is asymmetric with respect to the GPI allelic fre-
quencies: the frequencies of walkeri are mostly higher than
the corresponding S frequencies, whereas the frequencies
of barbouri are generally lower than the corresponding F fre-
quencies (Fig. 3). This discrepancy may indicate genetic
dominance of the walkeri genotype over the barbouri

69% selfing

FS

Fig. 2. Trivariate plot of the three GP! genotypes of Liguus fasciatus
in sections along a transect through Pinecrest hammocks 16 and 16a.
The upper curve represents the expected values of Hardy-Weinberg
equilibrium without self-fertilization, and the lower curve represents
the expected values with 69% self-fertilization. The numbered circles
indicate the genotypic combinations of the sections of the transect.
The location of section 10 (fixation of the FF genotype) is also the
location of sections 12-14.

genotype.

Roth and Bogan (1984) devised a system for describ-
ing phenotypic variation in Liguus fasciatus that incorporated
twelve distinct characters, each with two to four states. They
stated that they chose characters ‘‘..in which the alternate
states can be seen to segregate in randomly selected
material’ (Roth and Bogan, 1984). Under the Roth and Bogan
system, the three phenotypes present in PC 16 and PC 16a

HILLIS: PARTIAL SELF-FERTILIZATION OF TREE SNAILS 11

Phenotype

O 05 1.0
GPI allele

Fig. 3. Correlation of shell phenotypes and GPI alleles through
Pinecrest hammocks 16 and 16a. The open circles represent frequen-
cies of the walkeri phenotype and the S allele, and the closed circles
represent frequencies of the barbouri phenotype and the F allele.

are designated as follows: aurantius: CYBYS + EYU-M-LOPOA-
O-W-G*; barbouri: CYB8YS + EBU-M + LBPBA‘O-W+tG +; and
walkeri: CVBBYS-EBU-M + LBPBA+O+W+G+. As only these
three phenotypic combinations were observed among over
1,000 examined shells, the independence of the 12 characters
seems highly doubtful. If the 12 characters were independent,
one would expect 1024 phenotypic combinations of L.
fasciatus in PC 16-16a, rather than the observed three com-
binations. Instead, these phenotypes seem to be inherited as
single genes. This does not preclude the possibility of a few
tightly linked loci, however. Some of the 1021 unobserved
phenotypes do occur in other areas (Roth and Bogan, 1984),
but probably represent distinct alleles rather than recombina-
tions of the alleles present in PC 16-16a. Obviously, future at-
tempts at understanding the genetics of L. fasciatus shell
phenotypes must take into account self-fertilization.
Although partial self-fertilization of Liguus fasciatus is
sufficient to account for the high among-population variation
and low within-population variation observed throughout the
range of this species, this phenomenon does not account for
the overall low allozymic variability (Hillis et a/., 1987) com-
pared to the high morphological variability (Pilsbry, 1912, 1946)
found in Floridian Liguus. The low allozymic variaton could
be a result of a relatively recent invasion of few individuals
from Cuba, thus giving rise to fixation at most allozyme loci
through the founder effect. Fixation at most allozyme loci has
occurred in several introduced mollusks that are capable of
self-fertilization (Selander and Kaufmanm, 1973; McKracken
and Selander, 1980; Hillis and Patton, 1982). However, this
does not account for the high morphological variation seen
in Floridian populations of L. fasciatus. One possibility is that

the shell phenotypes are adapted to different local conditions.
However, adaptation is unnecessary to explain the distribu-
tion and variation of shell phenotypes. Instead, it is likely that
the genes responsible for shell phenotype undergo much
higher rates of mutation than do the allozyme loci, in which
case the partial self-fertilization of L. fasciatus would explain
the fixation of many of these phenotypes in the numerous
isolated Floridian populations of this species.

ACKNOWLEDGMENTS

| thank Oren Bass, Michael Dixon, Karen Ercolino, Archie
Jones, Lourdes Rodriguez, Modesto Sanchez, and Eric Zimmerman
for advise and assistance with this study, and Everglades National
Park and Big Cypress National Preserve for collecting permits. This
study was supported in part by a Nongame Wildlife Program Grant
from the Florida Game and Freshwater Fish Commission and by Na-
tional Science Foundation Grant BSR 8657640.

LITERATURE CITED

Brown, C. A. 1978. Demography, dispersal, and microdistribution of
a population of the Florida tree snail, Liguus fasciatus. Master’s
Thesis, University of Florida, Gainesville. 135 pp.

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Date of manuscript acceptance: 16 September 1988

MECHANICAL WEAR OF RADULAR DENTICLE CAPS
OF ACANTHOPLEURA GRANULATA (GMELIN, 1791)
(POLYPLACOPHORA: CHITONIDAE)

ROBERT C. BULLOCK
DEPARTMENT OF ZOOLOGY
UNIVERSITY OF RHODE ISLAND
KINGSTON, RHODE ISLAND 02881, U. S. A.

ABSTRACT

Mechanical wear of radular denticle caps of Acanthopleura granulata (Gmelin) was examined
using light and scanning electron microscopy. Between 6 and 11 transverse rows of teeth are involved
in feeding. Wear is first evident as slight abrasion and chipping in rows 8 to 11. Increased wear, chip-
ping, and occasional breakage of the cap subsequently occur. The conspicuous distal black tab on
the anterior surface, which is contiguous with the magnetite material of the posterior surface, begins
to wear quickly and usually disappears by row 6. The brown and yellow lepidocrocite region, in which
the tab is embedded, is mostly worn away by row 4, leaving only an amber base of apatite material
with an anterior surface of magnetite. The wearing cap is self-sharpening due to the differential hard-
ness of the leading surface of magnetite and the softer materials of the anterior surface. Presence
in the magnetite layer of fibers oriented at about 90° to the posterior surface also contributes to the
self-sharpening aspect. The fibers appeared to stop short of the posterior surface; a 90° ventral turn
near this surface was not observed. Progressive mechanical wear produces a chisel-shaped tooth that

provides an effective grazing capability and indicates a multi-tool feeding strategy.

Although functional morphology of the gastropod
radula has been the subject of several studies, little attention
has been paid to functional aspects of the highly complex
polyplacophoran radula. The relatively few studies of chiton
radulae have indicated that as the teeth are used they become
worn and are replaced. The process of mechanical wear of
the radula has not been described.

The polyplacophoran radula is a ribbon of teeth that
can reach a length more than half the length of the animal.
Chitons typically have 17 teeth per transverse row and many
rows of teeth exist (Fig. 1). Due to substratum contact, the
anterior-most teeth are subjected to great stress; Hickman
(1980) summarized these forces in her notable précis of
gastropod radula functional morphology. The two major lateral
teeth per row, also called the dominant teeth because of their
functional and visual prominence, are responsible for
substratum removal (Fretter and Graham, 1962; Steneck and
Watling, 1982; Bullock, 1986). In addition to its use in graz-
ing food particles, the radula appears responsible for crea-
tion of the protective homing scars of Acanthopleura gemmata
(Blainville, 1825) [Chelazzi and Focardi (1983); Chelazzi et al.
(1983)], Ceratozona angusta Thiele, 1909 (Schmidt-Effing,
1980), and Sypharochiton pelliserpentis (Quoy and Gaimard,

1835) (Boyle, 1970).

The formative end of the radula is located posteriorly,
and increasingly mature teeth are apparent anteriorly. As the
teeth at the anterior end of the radula become worn, they are
sloughed off and the radular ribbon advances to move fully
mature teeth into the feeding position.

Each major lateral tooth possesses a distinct cusp
heavily mineralized with the iron compound magnetite. The
first study of the mineralization process was published by Towe
and Lowenstam (1967), and other recent papers on the sub-
ject have appeared. Kim et al. (1986a) studied Clavarizona hir-
tosa (Blainville, 1825) and noted four developmental stages
of the radular ribbon: stage I, immature teeth composed of
a white organic matrix; stage II, with reddish brown denticle
caps; stage III, black magnetite becomes evident; and stage
IV, fully mineralized denticle caps. While this continuum en-
compasses the intriguing mineralization process, a complete
characterization of radular form must include a functionally
Critical fifth stage; the anterior-most teeth are being used and
becoming worn and lost due to the feeding process.

The structure and general composition of the
polyplacophoran radular denticle cap has been the subject
of various studies. The posterior surface, which in the feeding

American Malacological Bulletin, Vol. 7(1) (1989):13-19

13

14 AMER. MALAC. BULL. 7(1) (1989)

Fig. 1. Scanning electron micrograph of a portion of the radular rib-
bon of Acanthopleura granulata: dorsal view, anterior end toward top
of page; specimen from Las Tejitas, Isla de Margarita, Venezuela;
c, central tooth; cl, centro-lateral tooth; dc, denticle cap of major lateral
tooth; m, marginal teeth; mu, major uncinus; s, shaft of major lateral
tooth; w, wing of major lateral tooth (which is broken off as soon as
the major lateral tooth is moved into the feeding position); bar = 100
pm.

position is the scraping surface, is covered totally by a
substantial shield of magnetite. Colorful microarchitectural
units on the anterior surface provide species-specific dif-
ferences (Fig. 2). A marginal border of black magnetite sur-
rounds areas of brown and yellow lepidocrocite; ventral to the
lepidocrocite region is a more transparent, amber portion com-
posed of an apatite mineral (Lowenstam, 1967). A conspicuous
black tab of magnetite occurs distally in the lepidocrocite area
(Figs. 2A, 16) in most chitonid species.

Several authors have noted that the denticle caps
become worn and broken with use (Towe and Lowenstam,
1967; Mizota and Maeda, 1985; Lowenstam and Weiner, 1985;
Kim et a/., 1986a, b; van der Wal et a/., 1987), yet a descrip-
tion of this mechanical wear is lacking. | present information
in this paper about mechanical wear of the denticle cap of
the major lateral tooth of one species, Acanthopleura granulata
(Gmelin, 1791), that occurs abundantly from the Florida Keys
to the West Indies. Evidence from gastropod studies provides
valuable insight into the interpretation of the limited informa-
tion now available on the chiton radula.

Fig. 2. Distribution of microarchitectural units of the Acanthopleura
granulata denticle cap [adapted from Lowenstam (1967)]: A, anterior
view; B, posterior view; C, longitudinal section near tab; D. longitudinal
section through tab [a, amber component (apatite); b, brown com-
ponent (lepidocrocite); |, lepidocrocite; m, magnetite component
(black); t, tab (magnetite); y, yellow component (lepidocrocite)].

MATERIALS AND METHODS

Radulae of Acanthopleura granulata were examined
utilizing light and scanning electron microscopy (SEM).
Specimens were collected at northeastern Key Largo, Florida
(1986, n=13), Indian Key Fill, Florida (1977, n=44), Las Tejitas,
Isla de Margarita, Venezuela (C. Franz, leg. 1987, n=12), and
Playa Picua, La Blanquilla, Venezuela (C. Franz, leg. 1986,
n=8). Radulae were extracted from the animals, cleaned in
a heated 2N KOH solution, and placed in a series of distilled
water rinses in an ultrasonic cleaner. Some specimens were
kept in 70% ethanol and studied using a Wild M-8 stereo-
zoom dissecting microscope with a 1.6X adapter. Fully
mineralized denticle caps often cracked longitudinally when
air dried; however, the denticle caps of a few radulae were
broken with microforceps to create additional fracture sur-
faces. The radulae used for SEM were teased into pieces and
mounted on aluminum specimen mounts with Scotch Double-
Coated Tape No. 666. The specimens were coated with car-
bon and then 60% gold:40% palladium in a Denton DV-502
vacuum evaporator with a rotating/tilt device. All SEM work
was done on an ISI MSM-3 located in the Department of
Zoology, University of Rhode Island.

Characterization of tooth wear began by numbering the
transverse tooth rows beginning at the anterior end of the
radular ribbon. To measure denticle cap height, the anterior-
most 15 transverse rows of teeth were isolated and the in-
dividual major lateral teeth of one side were teased apart and
transferred to a double-coated tape surface or modeling clay
where they were positioned using an insect pin. Height
measurements were made using the Wild M-8 with a draw-
ing tube and stage micrometer. The height of unused denti-
cle caps was determined, and all worn caps were recorded
as a percent of this value.

Living Acanthopleura granulata from Crawl Key, Florida
(1987, n=7) were maintained in an aquarium. The feeding pro-
cess was observed and photographed using video equipment.

It was evident that two potential directional problems
exist. First, investigators of the mineralization process
understandably number the transverse tooth rows beginning
at the posterior formative end, although there is some
disagreement about where to begin numbering. However,
because tooth developmental processes are not always syn-
chronous among individuals (Lowenstam and Weiner, 1985),
and because an investigation of tooth wear involves only a
limited number of transverse tooth rows in the feeding posi-
tion at the anterior end of the radular ribbon, it appeared
crucial for mechanical wear studies that the numbering begin
with the most worn row at the anterior end of the ribbon and
proceed posteriorly. Only this procedure allowed an adequate
comparison among individuals.

A second directional problem could arise regarding the
surfaces of the denticle cap. The solid magnetite scraping
surface of the denticle cap faces posteriorly in the formative
and fully mature stages. Authors are consistent in calling this
surface the posterior surface. However, in an incorrect applica-
tion of a generalized gastropod model to the function of the
chiton radula, the radula is protruded from the mouth and the

BULLOCK: WEAR OF ACANTHOPLEURA RADULA 15

posterior surfaces of the teeth, which are now pointing
anteriorly, are moved anteriorly and then dorsally, bringing
grazed particles into the buccal cavity. It would be tempting
to stress the functional process and to designate the scrap-
ing surface as the anterior surface. Although this generalized
model might be seen in some chiton groups, literature reports
indicate that, at least in the Suborder Chitonina, the oppos-
ing major lateral teeth, that spread apart as the chiton begins
to feed, converge medially, forming characteristic grazing
marks that are perpendicular, not parallel, to the longitudinal
axis of the animal (JUch and Boekschoten, 1980; Bullock,
1986). It seems best to retain the current terminology and ac-
cept the fact that the chiton produces and manipulates the
radular ribbon in a way that greatly changes these directions
during the feeding process.

RESULTS

Mechanical wear of the denticle caps of the major
lateral teeth was evident in all individuals examined. This wear
was seen as abrasion, slight chipping, and, occasionally,
breakage. Observation of Acanthopleura granulata feeding on
the sides of glass-walled aquaria indicates that in each feeding
event about 7 to 10 pairs of denticle caps typically sweep the
substratum. It is unclear exactly how many of these teeth ac-
tually make contact with the substratum, but studies utilizing
Plexiglas indicate that each feeding event results in 3 to 6
pairs of grazing marks. Irregular substrata could provide quite
different results. Abrasion and reduction of denticle cap height
begin soon after the teeth move anteriorly enough to be in-
volved in the feeding process (Fig. 3). Use of the teeth causes
sporadic chipping of the distal end of the denticle caps, and
plots of denticle cap height are irregular because of this
phenomenon. Denticle cap height declines with use until the
major lateral tooth is discarded. Occasional denticle caps are
broken off near their base.

The distribution of the different components of the den-
ticle cap was evident by the color pattern on the anterior sur-
face. The colors of the non-magnetite units were not the same

Denticle Cap Height (%)

12 1110 9 8 7

6 5 4 3 2 1
Tooth Row (Posterior ——> Anterior)

Fig. 3. Denticle cap height vs. transverse tooth row at anterior end
of radular ribbon; Las Tejitas, Isla de Margarita, Venezuela, n = 6.

Figs. 4-15. Outline drawings of denticle caps showing mechanical
wear, anterior and lateral views (single individual, NE Key Largo,
Florida). Figs. 4, 5. Transverse tooth row 14 (unused), with all ex-
terior components fully visible. Figs. 6, 7. Row 11, slight chipping of
distal edge evident. Figs. 8, 9. Row 6, increased chipping, decreased
height, reduction of distal tab, magnetite, and lepidocrocite units. Figs.
10, 11. Row 5, tab nearly gone (typically lost by this time), lepidocrocite
worn away except at lateral margin. Figs. 12, 13. Row 3, tab absent,
mostly amber base with magnetite scraping surface. Figs. 14, 15.
Row 1, lepidocrocite absent. Anterior views do not differentiate the
brown and yellow bands of lepidocrocite; lateral view only shows
magnetite unit; bar = 200 um.

as Lowenstam (1967) reported for Acanthopleura echinata. The
lepidocrocite in A. granulata was seen as a distal brown band
and a more ventral yellow layer (Fig. 2A). Lowenstam (1967)
noted that the colors of this unit differ due to transparency
and their proximity to other components.

The denticle caps of Acanthopleura fixed and preserved
in 70% ethanol tended to separate quite easily from the shaft
of the major lateral tooth, especially after a few years in
alcohol. This situation was not seen in freshly preserved
animals, and it is obvious that in living Acanthopleura the den-
ticle cap is very securely attached to the shaft.

Light and scanning electron microscopy of denticle
caps allowed a clear view of mechanical wear as well as in-
formation about microstructure. Fully mineralized caps often
fractured when air dried, and the resulting cracks afforded
various sectional views of cap morphology. The microarchitec-
tural units were easily seen with light microscopy because
of their color differences (Figs. 4-15). Tooth rows 8 to 11 showed
slight abrasion and chipping of the black magnetite of the
distal end. By row 5, the black tab had disappeared totally
in most cases. Loss of most of the lepidocrocite layer was evi-
dent by rows 4 to 5 (Fig. 10). The denticle caps in rows 1 to
4 were quite stubby and only the amber apatite base and the
black magnetite layer of the posterior (cutting) surface were
present (Figs. 12-15).

Abrasion of the anterior surface was readily apparent
with scanning electron microscopy (Figs. 16-21). The wear-
ing teeth remained chisel-shaped. Examination of fracture sur-

16 AMER. MALAC. BULL. 7(1) (1989)

Figs. 16-21. Scanning electron micrographs of Acanthopleura granulata denticle caps. Fig. 16. Unused denticle cap showing rounded distal
end and granular tab; Indian Key Fill, Florida; bar = 100 um. Fig. 17. Lateral view of denticle cap from first (most worn) transverse row; note
abrasion on sides and anterior surface; Las Tejitas, Isla de Margarita, Venezuela; bar = 100 um. Figs. 18, 19. Distal lip of worn caps showing
abrasion and self-sharpening due to orientation of fibers in magnetite unit. Fig. 18. Tooth row 2, bar = 10 um. Fig. 19. Tooth row 4, bar =
20 um; Las Tejitas, Isla de Margarita, Venezuela. Fig. 20. Cross section through distal portion of denticle cap showing granular tab of anterior
surface (upper left) and the magnetite fibers that extend from tab toward posterior surface (lower right); NE Key Largo, Florida; bar = 10 um.
Fig. 21. Fractured distal portion of denticle cap revealing fibrous microstructure of posterior surface; note smooth margin at posterior surface

(lower left); NE Key Largo, Florida; bar = 20 um.

BULLOCK: WEAR OF ACANTHOPLEURA RADULA 17

faces revealed the fibrous construction of the magnetite layer
(Fig. 21). These fibers, that at times appear as lamellar
clusters, are oriented at about 90° to the posterior surface of
the denticle cap. | was unable to verify that the fibers bend
ventrally at or near the posterior surface; a smooth margin
was observed in this region (Fig. 21).

Mechanical wear of the radula apparently proceeds at
different rates in different geographic locations. Wear quick-
ly reduces denticle cap height past the black tab in some
populations, but in other localities the tab is visible through
more tooth rows. In severe cases, the first 4 or 5 tooth rows
are missing all of the tab and all lepidocrocite. Even in a single
population, there is considerable wear difference between
individuals.

DISCUSSION

The presence of iron compounds in the denticle cap
material of limpets and chitons has intrigued biologists and
geologists for decades. The hardened nature of the radula
has important implications for those interested in the biological
precipitation of these iron compounds and the behavioral and
ecological consequences of this hardness. Pioneering work
in the area of limpet radular structure was done by Jones et
al. (1935) and Runham and his co-workers (Runham and
Thornton, 1967; Runham et a/., 1969). A few investigators have
reported on the radula of the Polyplacophora (Tomlinson,
1959; Runham, 1963; Carefoot, 1965; Lowenstam, 1967; van
der Wal et al., 1987), and recently some workers have focused
their efforts on the biomineralization process (Towe and
Lowenstam, 1967; Lowenstam and Weiner, 1985; Mizota and
Maeda, 1985; Kim et a/., 1986a, b). These studies have shown
that the cusps of limpets and chitons are a composite of dif-
ferent materials and that the specific materials present and
their distribution have important functional aspects.

Lowenstam (1967) presented diagrams of the posterior,
anterior, and longitudinal cross section of the denticle cap of
Acanthopleura echinata (Barnes, 1824). | accept these
diagrams as factually correct with the exception of the
longitudinal cross section (similar to my Fig. 2C) which fails
to show the existence of the conspicuous black tab of the
distal anterior surface. Many of the longitudinal fractures that
occur during drying fail to develop at the site of the black tab.
However, | observed fracture surfaces in the region of the
tab, and | was able to see that the magnetite at this site is
broad at the surface but it narrows before joining the magnetite
of the posterior surface. Of course, examination of the anterior
surface of a denticle cap, including Lowenstam’s diagram,
would dictate the inclusion of the tab in a longitudinal cross
section (Fig. 2D). Lowenstam (1967) stated that due to the in-
tervening lepidocrocite, the magnetite layer does not directly
contact the apatite. However, | observed that the tab magnetite
penetrates the apatite at its most distal portion (Fig. 20), and
| saw no lepidocrocite where this intrusion occurs. The con-
tinuity of the magnetite between the tab and the posterior sur-
face provides substantial support for the tab (Fig. 2D).

The magnetite is present in functionally important
areas of the Acanthopleura and Chiton denticle cap. The en-

tire posterior surface, the scraping surface, is completely
covered with a substantial layer of magnetite. The anterior sur-
face, which is less subjected to the rigors of substratum con-
tact, has a narrow marginal band of magnetite that is con-
tiguous with that of the posterior surface. Elsewhere across
the distal anterior surface, only lepidocrocite is present ex-
cept for the magnetite of the tab. Bullock (1986) stated that
use of magnetite in the denticle cap is conserved. However,
it is important to recognize that the denticle caps function very
effectively and that any additional magnetite might provide
additional hardness but disrupt the self-sharpening
phenomenon. For example, more magnetite on the anterior
surface would provide protection but would interfere with self
sharpening. Presence of the tab provides some of this pro-
tection but at the same time allows the softer lepidocrocite
and apatite layers to become worn.

| suggested previously (Bullock, 1986) that the tab
could protect the otherwise unprotected anterior surface dur-
ing movement of the radula across the substratum. The tab
often begins to disappear quickly when the cusp is used for
feeding. At this point, many of the cusps involved in feeding
already have worn down past the tab. Any protection provid-
ed by the tab during feeding is better than no protection, and
the chiton is well served by even a brief existence of the tab.
However, the observation that the tab disappears quickly in-
dicates that the tab could also function to protect the anterior
surface prior to tooth use. The radular ribbon remains inwardly
curled until the tooth rows are moved into the feeding
position; some aspects of radular morphology reflect passive
accommodation of the teeth because of ‘‘curled’’ contact;
other morphological features appear to have a pre-feeding
functional basis. Lacking protection, the denticle caps could
abrade each other during normal movement of the animal over
irregular substrata. Within column abrasion of the anterior sur-
face by the posterior surface of magnetite of the next cap is
prevented or minimized by: (1) interleaving of the major
uncinus (Fig. 1, mu) between denticle caps, and (2) presence
of the tab. Contact with the opposing denticle cap is prevented
by the wing of each major lateral tooth (Fig. 1, w). The wings
are broken off immediately upon movement of the tooth row
into the feeding position.

The existence of a fibrous microstructure in the
magnetite was not unexpected. Runham and Thornton (1967)
and Runham et al. (1969) noted 800 A thick fibers in the
mineralized cusps of Patella vulgata Linnaeus, and Towe and
Lowenstam (1967) had reported a fibrous network in the
development of the denticle cap of Cryptochiton stelleri (Mid-
dendorff, 1847). More recently, van der Wal et a/. (1987) brief-
ly commented on ‘‘closely packed rod-shaped and elongate
concavo-convex (trough-shaped) units’’ in the denticle cap
of Chiton olivaceus Spengler, 1797. According to the latter
authors, within the magnetite unit the fibers are oriented
perpendicular to the posterior surface, but they turn ventral-
ly 90° near the posterior surface. Although fibers and lamellar
clusters of fibers are easily observed in SEM of fractured
Acanthopleura granulata denticle caps, | was unable to see
clear evidence of their ventral turn in my preparations. | almost
always found that the visible fibers stopped short of the

18 AMER. MALAC. BULL. 7(1) (1989)

posterior edge, which appeared rather smooth in cross sec-
tion. This smooth boundary could be the region where the
fibers are oriented parallel to the posterior surface, but my
preparations, and perhaps the limited resolution of the
available SEM, did not document any change in fiber
orientation.

The existence of fibers within a matrix is a critical
feature of the functional morphology of the polyplacophoran
denticle cap, but just how these components provide the ob-
vious hardness of the tooth is a question with an exceeding-
ly elusive answer. Vincent (1980: 132) concluded that ‘‘the
hardness of biological materials has not attracted the atten-
tion of experimentalists . . . It is not only a difficult measure-
ment to make but its interpretation can be fiendishly difficult.
The hardness of artificial composites, whose variables can
be closely controlled, is a subject that has hardly yet been
broached, let alone the hardness of the much more complex
biological composites.’

The mineralized cusps of limpets and chitons wear
down yet maintain a rather sharp cutting edge (Runham and
Thornton, 1967; Runham et a/., 1969; Bullock, 1986; van der
Wal et al., 1987). This self-sharpening phenomenon is due
to the hard but thin leading posterior surface and the relative
softness of the much thicker anterior portion of the cusp.
Furthermore, the orientation of the fibers assists by fractur-
ing lengthwise, which helps to maintain the wedge angle. The
wedge angle of Acanthopleura granulata is about 60° in
unused teeth, but with wear this angle can increase to as
much as 70°. The fibers parallel to the surface along the
leading posterior edge, which are well documented in Patella
and reported in Chiton olivaceus (van der Wal et al., 1987),
are less susceptible to wear (Runham et al., 1969), and this
distal edge is the actual cutting portion of the denticle cap.
As denticle caps of A. granulata become worn with use, the
distal edge is seen as a slightly thickened lip (Figs. 17-19).

The fibers of the tab magnetite that proceed to the
posterior surface are oriented to assist, or at least allow, self-
sharpening when mechanical wear affects this level; this
orientation also means that the fibers at the anterior surface
are nearly perpendicular to it, and the exterior portion of the
tab has increased resistance to wear. The conspicuous
granules of the tab are surface protrusions of the underlying
fibers or clusters of fibers of magnetite (Fig. 20).

The form of the unused denticle cap could perhaps
not be the most efficient morphology; as Hickman (1980)
noted, it could be important functionally for the teeth to wear
down to be efficient. A variation of this theme would be the
recognition that in the feeding position, the continuum from
new to worn teeth would provide different capabilities and that
it would be advantageous to graze on heterogeneous
substrata with a multi-tool approach. All denticle caps of this
continuum, including those of the anterior-most transverse
row, suffer mechanical wear, indicating that the major lateral
teeth continue to function until they are lost.

No studies of chiton radulae have examined intra-
specific differences due to life on substrata of varying hard-
ness. The number of teeth that become substantially worn
varied greatly from individual to individual within a single

population, and the anterior-most teeth were not worn to the
same degree. In some Acanthopleura granulata maintained
in aglass aquarium for two weeks, most of the denticle caps
in the feeding position still retained at least some of the distal
anterior tab, indicating that wear had not proceeded at the
same rate as that of the teeth from field-collected and fixed
individuals.

ACKNOWLEDGMENTS

This paper was presented as part of a symposium on the
biology of the Polyplacophora at the 1987 annual meeting of the
American Malacological Union held at Key West, Florida. Some of
my comments reflect the results of discussions with participants in
this symposium. Freshly preserved examples of Acanthopleura
granulata from Venezuela were provided by C. Franz of the Univer-
sity of Rhode Island. Dr. R. Turner of Harvard University assisted with
the collection of living A. granulata from the Florida Keys and offered
information that allowed the transportation of these specimens. Ad-
ditional specimens from the Florida Keys were obtained during
fieldwork sponsored by the College of Arts and Sciences and a Grant-
in-Aid-of-Research from the Office of the Coordinator of Research
at the University of Rhode Island. | thank J. S. Cobb of the Depart-
ment of Zoology for allowing the use of video equipment, and P.
Langer of Gwynedd-Mercy College for providing some reference
materials. | appreciate constructive comments on the manuscript pro-
vided by C. Franz, C. R. Shoop, R. Turner, and two anonymous
reviewers.

LITERATURE CITED

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pelliserpentis (Mollusca: Polyplacophora). New Zealand Jour-
nal of Marine and Freshwater Research 4:364-384.

Bullock, R. 1986. Functional morphology of some chitonid radulae
(Polyplacophora: Chitonidae). American Malacological Bulletin
4:114-115.

Carefoot, T. 1965. Magnetite in the radula of the Polyplacophora. Pro-
ceedings of the Malacological Society of London 36:203-212,
pl. 10.

Chelazzi, G. and S. Focardi. 1983. Different space exploitation in two
sympatric intertidal chitons. Monitore Zoologico Italiano (n.s.)
17:185-186.

Chelazzi, G., S. Focardi, J. L. Deneubourg and R. Innocenti. 1983.
Competition for the home and aggressive behaviour in the
chiton Acanthopleura gemmata (Blainville) (Mollusca:
Polyplacophora). Behavioral Ecology and Sociobiology 14:15-20.

Fretter, V. and A. Graham. 1962. British Prosobranch Molluscs. Ray
Society, London. 755 pp.

Hickman, C. 1980. Gastropod radulae and the assessment of form
in evolutionary paleontology. Paleobiology 6:276-294.
Jones, E., R. McCance and L. Shackleton. 1935. The role of iron and
silica in the structure of the radular teeth of certain marine

molluscs. Journal of Experimental Biology 12:59-64, pl. |.

Juch, P. and G. Boekschoten. 1980. Trace fossils and grazing traces
produced by Littorina and Lepidochitona, Dutch Wadden Sea.
Geologie en Mijnbouw 59:33-42.

Kim, K.-S., J. Webb, D. Macey and D. Cohen. 1986a. Compositional
changes during biomineralization of the radula of the chiton
Clavarizona hirtosa. Journal of Inorganic Biochemistry
28:337-345.

BULLOCK: WEAR OF ACANTHOPLEURA RADULA 19

Kim, K.-S., J. Webb and D. Macey. 1986b. Properties and role of fer-
ritin in the hemolymph of the chiton Clavarizona_hirtosa.
Biochimica et Biophysica Acta 884:387-394.

Lowenstam, H. 1967. Lepidocrocite, an apatite mineral, and magnetite
in teeth of chitons (Polyplacophora). Science 156:1373-1375.

Lowenstam, H. and S. Weiner. 1985. Transformation of amorphous
calcium phosphate to crystalline dahllite in the radular teeth
of chitons. Science 227:51-53.

Mizota, M. and Y. Maeda. 1985. Magnetite in chitons. Journal of the
Physical Society of Japan 54:4103-4106.

Runham, N. 1963. The histochemistry of the radulas of Acanthochitona
communis, Lymnaea stagnalis, Helix pomatia, Scaphander
lignarius, and Archidoris pseudoargus. Annales d’Histochimie
8:433-441.

Runham, N. and P. Thornton. 1967. Mechanical wear of the gastropod
radula: a scanning electron microscope study. Journal of
Zoology, London, 153:445-452.

Runham, N., P. Thornton, D. Shaw and R. Wayte. 1969. The
mineralization and hardness of the radular teeth of the limpet
Patella vulgata L. Zeitschrift fur Zellforschung 99:608-626.

Schmidt-Effing, U. 1980. Beobachtungen zum Heimfinde-Verhalten
und zum Aktivitats-Rhythmus von Chiton stokesii Broderip,
1832 mit Anmerkungen auch zu /schnochiton dispar (Sower-
by, 1832) (Polyplacophora). Zoologischer Anzeiger, Jena,
205:309-317.

Steneck, R. and L. Watling. 1982. Feeding capabilities and limita-
tion of herbivorous molluscs: a functional group approach.
Marine Biology 68:299-319.

Tomlinson, J. 1959. Magnetic properties of chiton radulae. Veliger 2:36.

Towe, K. and H. Lowenstam. 1967. Ultrastructure and development
of iron mineralization in the radular teeth of Cryptochiton stelleri
(Mollusca). Journal of Ultrastructural Research 17:1-13.

van der Wal, P., J. Videler, PR. Havinga and R. Pel. 1987. Architecture
and chemical composition of the magnetite-bearing layer in
the radula teeth of Chiton olivaceus (Polyplacophora).
Ultramicroscopy 21:204-205.

Vincent, J. 1980. The hardness of the tooth of Patella vulgata L. radula:
a reappraisal. Journal of Molluscan Studies 46:129-133.

Date of manuscript acceptance: 27 April 1988

BEHAVIOR, BODY PATTERNING, GROWTH AND LIFE HISTORY OF

OCTOPUS BRIAREUS CULTURED IN THE LABORATORY

ROGER T. HANLON
MARTIN R. WOLTERDING
THE MARINE BIOMEDICAL INSTITUTE
THE UNIVERSITY OF TEXAS MEDICAL BRANCH
GALVESTON, TEXAS 77550-2772, U.S.A.

ABSTRACT

A total of 10 years of laboratory observations are reported, centering mainly upon four groups
of Octopus briareus Robson cultured through the life cycle. Details are given for rearing methodologies,
system design, egg development, hatching and feeding. A detailed analysis of growth indicated that
this species grows at a fast exponential rate (4.8% mean increase in body weight per day) for the first
18 - 20 weeks, then growth slows to a logarithmic rate for the remaining 30 weeks of the life span.
Growth is allometric, with arms III growing faster and larger than arms |, Il and IV to become a chief
morphological character of the species. O. briareus is susceptible to bacterial skin lesions and this
species is strongly cannibalistic. Life span is estimated to be 10 to 17 months.

The emphasis of the study was behavior. The morphology and development of body patterns
are described, with detailed descriptions of the 18 chromatic, four textural, nine postural and four
locomotor components of patterning. Aspects of exploratory behavior as well as intraspecific (agonistic,
reproductive) and interspecific interactions (attack, defense) are described using body patterns as the

bases of description.

Octopuses have been the subject of folklore for cen-
turies (Lee, 1875; Aristotle In: Peck, 1970; Lane, 1974), but
more recent and comprehensive scientific accounts of octopus
biology (e.g. Robson, 1929a, b, 1932; Pickford, 1945; Wells,
1978; Boyle, 1983, 1987) have helped clarify our understand-
ing of these active carnivores that have a short life span but
occupy a high trophic level in the marine ecosystem. Many
gaps remain, especially in our knowledge of how octopuses
behave, grow and reproduce in nature. This study was initiated
in 1969 to fill in some of those gaps for Octopus briareus
Robson.

This laboratory study was conducted by Wolterding in
Miami from 1969-1971, by Hanlon in Miami from 1972-1975,
and later by Hanlon and Forsythe in Galveston from
1980-1984. In all, four groups of Octopus briareus were
cultured through the life cycle so that the observations herein
come from hundreds of animals from different populations
and year classes.

LABORATORY CULTURE

MATERIALS, MEASUREMENTS AND WATER
QUALITY

In 1969 and 1972, Octopus briareus eggs or gravid

females were obtained by skin or SCUBA diving in shallow
water (1 - 2 m) south of Miami, Florida (Card Sound, Soldier
Key, Key Largo) and transferred to the open seawater system
at the Rosenstiel School of Marine and Atmospheric Science,
University of Miami, on Virginia Key. The system was de-
scribed in detail by Myrberg (1969). All tanks were subject to
indirect sunlight. Therefore the light, temperature, and salinity
cycle closely resembled the natural environment throughout
the year. Wolterding reared five adults to maturity (one laid
fertile eggs) from about one hundred hatchlings in 1969-1971
and made behavioral observations (between 1000 and 1200
hours) on 24 individual octopuses, some of which were field-
caught and maintained. Hanlon reared eight adults to maturity
from about one hundred hatchlings in 1973 and made growth
measurements and behavioral observations.

In 1981 in Galveston, Texas, 34 eggs were obtained
from the Dallas aquarium, where a female imported from
Florida had laid eggs. Eight octopuses were cultured through
the life cycle and produced second generation eggs and
hatchlings. In 1982, portions of two broods of eggs (n = 300)
were collected from Sweeting’s Pond on Eleuthera Island,
Bahamas, and shipped to Galveston. Thirty octopuses were
cultured through the life cycle and five females produced
second generation eggs. A growth analysis and extensive

American Malacological Bulletin, No. 7(1) (1989):21-45

21

22 AMER. MALAC. BULL. 7(1) (1989)

observations on body patterning were made on these oc-
topuses. The large closed seawater system in which they were
cultured has been described by Hanlon and Forsythe (1985).

Small octopuses were anaesthezied either in 12%
ethyl alcohol/sea water or 1/2 - 22% ethyl carbamate
(urethane)/sea water. For adults, the concentrations were
raised to 3% of each solution. It usually took one to four
minutes to completely relax the octopuses. Urethane relaxes
the arms very well and is excellent for growth measurements,
but the animals reacted violently to it for the first 30 seconds.
Alcohol does not relax the arms very well but the octopuses
react calmly to it. Therefore, in later experiments on adults,
a combination was used, consisting of either (1) 3% alcohol
followed by 11/2% urethane, or (2) 3% alcohol mixed with 172%
urethane. The animals did not react violently and the arms
relaxed well. It was important to wash the animal off briefly
with fresh sea water and to keep it slightly moist while measur-
ing it. Afterwards the octopus was held by hand and a mild
jet of sea water sprayed into the mantle over the gills. Usual-
ly in one to three minutes the octopus ventilated and regained
muscle control and alertness. No mortalities resulted from this
procedure.

Growth measurements were taken with dial calipers
on live, anaesthetized octopuses throughout the life cycle. The

E +—F G

posterior |
i

ventral

Fig. 1. Growth measurements and terminology. A - mantle length,
B - mantle width, C - total length, D - arm length. For behavioral ter-
minology: E - arms, F - head, G - mantle.

length measurements are illustrated in figure 1. First and third
right arm lengths were measured since they represented the
shortest and longest arms, respectively. In all cases the re-
laxed arms were gently stroked outward then measured
several seconds later after they were stationary. For wet
weights, octopuses were held momentarily with the mantle
highest so that excess water drained out, then weighed to the
nearest 1.0 mg.

Still photography was used extensively in document-
ing behavior and body patterns. Approximately 930
photographs were taken in the laboratory and 230 in the field.
A 35 mm Asahi Pentax or Nikon camera system was used
with various close-up lens and bellows units (for magnifica-
tion up to 16 : 1) and color slide film and electronic flash.
Underwater photographs were taken with a Nikon F and 55
mm Micro-Nikkor lens and electronic flash in a Lexan housing.

Water quality in the open system in Miami was
monitored only for temperature (mean 25°C, range 18 - 29°C)
and salinity (mean 33 ppt, range 27 - 36 ppt). The closed
systems in Galveston were monitored for many parameters.
Most octopuses were cultured between 20 and 26°C, 32 - 38
ppt salinity and at a pH of 7.8 - 8.2. A pH below 7.5 begins
to affect octopus metabolism adversely. Sustained levels of
ammonia-nitrogen (NH, -N) and nitrite-nitrogen (NOz2 -N) were
generally kept below 0.2 mg//, but on rare occasions oc-
topuses tolerated levels as high as 10.4 mg// NH, -N and 0.3
mg/! NOz -N for a few days. Sustained levels of nitrate-nitrogen
(NO3 -N) were generally kept below 200 mg//, but levels as
high as 650 mg/! for several days produced no observable
ill effects. The key to this tolerance of high levels of
nitrogenous waste was maintenance of pH between 7.8 and
8.2. These high levels of tolerance are in general agreement
with the findings of Hirayama’s (1966) short-term experiments
on Octopus vulgaris Cuvier in Japan. The 2600 / closed
seawater system in which O. briareus was Cultured in 1982
was found to support 15 kg of octopus biomass and still main-
tain nitrogenous levels at an acceptable level (Hanlon and
Forsythe, 1985).

REARING METHODS

In 1969 Wolterding reared hatchlings in floating plex-
iglass boxes (with screened sides) for the first 50 days, in 12
/ aquaria until day 102 and in 40 / glass aquaria thereafter.
Plexiglass sheets were affixed over the tops of the aquaria
to prevent escapes. For dens, small clear plastic vials or ar-
tificial grass were used for the youngest animals, while large
rocks, unglazed earthenware and flower pots were provided
for large octopuses. Hatchlings were hand-fed disarticulated
crab legs for the first two weeks; thereafter they ate small live
crabs (Uca sp.).

In 1973 Hanlon reared some hatchlings as Wolterding
did, while other octopuses were reared solely on live crabs
in a broad but shallow (7 cm water depth) water table in which
octopuses hiding in the bottom of the floating artificial grass
could see and easily attack the crabs crawling just below
them. Large numbers of crabs were kept in the tank to con-
centrate the food source and thus enhance feeding. As the

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 23

octopuses grew, an excess of small mollusc shells and ir-
regularly shaped rocks were added for shelter. The distance
from the water level to the top of the tank sides was 12 cm
and thus prevented small octopuses from crawling out. At two
months, octopuses were moved to 75 / aquaria with
flower pots to hide in, and hinged tops were secured to the
tanks or screened wooden frames were fit snugly over the
top of the tanks. For growth studies individual animals were
reared in separate aquaria.

In 1981 and 1983 in Galveston, the same group-rearing
concept of Hanlon’s 1973 study was used in which shallow
water tables were stocked with (1) octopuses (density
equivalent to 300 - 700 hatchlings per m2), (2) numerous small
polyvinyl chloride (PVC) sections of pipe for hiding, and (3)
high densities of small mysid shrimps (genus Mysidopsis).
The sizes of pipe and shrimp were increased as the octopuses
grew. Due to the cannibalistic nature of Octopus briareus, oc-
topuses had to be reared in separate containers after about
two months to achieve good survival (Hanlon and Forsythe,
1985). Octopuses up to 1 kg could be grown in containers
as small as 28 cm diameter and 21 cm deep with screened
sides to allow water exchange. For some growth data, some
octopuses were cultured through the entire life cycle in
perpetual isolation, beginning with 50 m/ plastic cups with
screened sides for hatchlings; these animals showed no ob-
vious physiological or behavioral deficits.

EGG DEVELOPMENT AND HATCHING

The large eggs measure 10 - 14 mm long and 4-5
mm wide, with a stalk 5 - 10 mm long. The stalks of seven
to 34 eggs (mean 25) were intertwined onto a central strand
8 - 10 cm long that was attached by the female to the
substratum (Fig. 2). Development took 50 - 80 days at 19 -
25°C, during which time the embryos underwent two rever-
sals (Fig. 3). From two to seven days before hatching the em-
bryo rotated 180 degrees toward the distal end of the egg and
a seam developed across the distal third of the egg. Physical
stimulation of the egg (by the mother or experimenter) resulted
in (1) increased respiration and arm movements, (2) expan-
sion of chromatophores over the embryo, (3) one or two rota-
tions of the embryo and (4) occasional squirting of ink within
the egg. This increased movement (and secretions from
Hoyle’s organ, which releases a lytic enzyme) caused the egg
seam to split, and hydrostatic pressure within the egg ejected
fluid and part of the mantle out of the egg (Fig. 4). Within
seconds or a few minutes the octopuses managed to jet or
squeeze their way out of the egg. Hatching occurred generally
at night, and hatching success was very high, mainly greater
than 95%. Females characteristically laid and actively brood-
ed (Fig. 5) about 200 - 500 eggs, although one laboratory-
cultured 500 g female in Galveston laid 955 eggs in 1981.
Many eggs in this study were removed from the mother and
brooded artificially by suspending them over gently bubbling
water. The hatchlings were fully formed, miniature adults that
had no planktonic phase; they would often crawl, ink, jet,
change color and feed within moments of hatching
(Messenger, 1963). They could survive up to ten days on in-
ternal yolk. Sometimes the hatchlings had a small external

yolk sac (1 - 3 mm); hatchlings seven to ten days premature
had a large external yolk sac and did not survive well.

FOODS

Live crabs are the favorite food of Octopus briareus of
all sizes, but they will accept a wide range of crustaceans,
molluscs, polychaetes and fishes. They will readily attack and
capture prey that are from 1/3 to 2X their own mantle length.
Table 1 lists species that O. briareus will attack and/or eat in
the laboratory. Over 500 feedings were observed and a few
observations are noteworthy. The gastropods Fasciolaria tulipa
Linné and Strombus gigas Linné were attacked repeatedly
but always released within one minute. The stomatopod
Gonodactylus sp. was eaten despite having stabbed the oc-
topus several times. Crabs were relatively defenseless and
appeared paralyzed within three minutes after capture. Her-
mit crabs were eaten by pressing the apertures of their shells
against the buccal mass of the octopus; after five to ten
minutes, one arm reached into the shell and extracted the
crab. The horseshoe crab, Limulus polyphemus Linné, ap-
peared to be too difficult to eat. It took about 30 minutes to
see the effect of paralysis from the octopus venom, and the
octopus tried unsuccessfully for five and one-half hours to
disarticulate and eat the crab.

GROWTH RATES BY LENGTH AND WEIGHT

The following growth data are available: five octopuses
reared to maturity in an open seawater system in Miami in
1969; eight animals grown to maturity in the same open
system in Miami in 1973; eight octopuses reared to maturity
in a closed seawater system in Galveston in 1981; and 20
animals weighed and measured from hatching to maturity dur-
ing a large-scale culture effort in a closed seawater system
in Galveston in 1983. In the latter two experiments, numerous
broods of second-generation eggs were produced. The growth
results in all experiments were remarkably consistent and in-
dicate clearly that Octopus briareus increases in length and
weight exponentially during the first 16 to 20 weeks of the life
cycle, after which growth slows to a more typical logarithmic
form until senescence and death. At hatching O. briareus is
approximately 6 mm mantle length (ML) and 95 mg wet weight
(WW). The largest laboratory reared animals were a female
of 175 mm ML and 1,055 g at 252 days and a male of 150
mm ML and 1,083 g at 324 days.

Mean length measurements at hatching were: 15.0 mm
total length (TL); 5.5 mm mantle length (ML); 5.0 mm mantle
width (MW); 8.0 mm first arm length (AL,); and 9.0 mm third
arm length (AL3). The measured dimensions are illustrated
in figure 1. First arm length represents the shortest arm length
and third arm length represents the longest because the arms
are in the order of 2=3.4.1. Mantle length is the standard
length measurement in most cephalopod studies. For the 20
animals reared through the life cycle in 1983, growth results
are given for ML (Fig. 6), MW (Fig. 7), TL (Fig. 8), AL, (Fig.
9) and AL; (Fig. 10). Regression analyses of length data in-
dicated in all cases that there was a distinct slowing in growth
in the period of 16 - 20 weeks. Collectively, the data were
best split at the 18-week period. A similar break was seen in

24 AMER. MALAC. BULL. 7(1) (1989)

Fig. 2. Freshly laid egg clusters entwined around a central stalk, and at bottom 50-day-old eggs suspended inside a flower pot. Fig. 3. Egg
development. From top to bottom: newly laid egg with embryo forming at right; octopus embryo before second inversion (45 days old); and
dorsal and ventral views of octopus after second inversion (50 days old). Fig. 4. Hatching sequence. From left to right: egg capsule is split
at posterior end; octopus squeezing its mantle out; only the arms remaining inside the egg capsule; newly hatched octopus. Fig. 5. Female

octopus brooding eggs in the protective posture ( Comp. 24).

analyses of the 1973 and 1981 data. Collectively, these length
measurements indicate growth rates of 1.5 to 1.9% increase
in body length per day during the exponential phase (d1- 126),
and progressively slower rates during the logarithmic phase
until senescence begins.

Analysis of wet weight increases in the 1975, 1981 and
1983 data (n = 41) revealed that growth was clearly exponen-
tial for the first 18 - 20 weeks under laboratory conditions (Fig.
11). Clearly, growth remains rapid over a long period. After

18 - 20 weeks, growth was best described by the logarithmic
equation form. Full details of the 1983 weight data are given
in table 2. Figure 12 illustrates changes in growth rates deter-
mined over short intervals. The highest growth rates occurred
between weeks four and eight and there was an inexplicable
cyclic pattern of increasing and decreasing rates during the
first eight weeks. Overall, the mean growth rate over the first
18 - 20 weeks was 4.8% increase in body weight per day,
which coincided well with the growth rate exponents of 4.6

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 25

Table 1. Prey organisms that Octopus briareus attacked or ate under laboratory conditions.

Actively Actively
Animal Attacked Eaten Animal Attacked Eaten
Polychaetes Crabs:
Chaetopterus variopedatus (Renier) No Yes Aratus pisonii (Milne-Edwards) Yes Yes
w/ tube Calappa flammea (Herbst) Yes Yes
Hermodice carunculata (Pallas) No Yes Callinectes ornatus Ordway Yes Yes
Onuphis magna Andrews wi/ tube Yes No C. sapidus Rathburn Yes Yes
O. magna wio tube Yes Yes Cardisoma guanhumi Latréille Yes Yes
Clibinarius vittatus (Bosc) Yes Yes
Molluscs Coenobita clypeatus (Herbst) Yes Yes
Bivalves: Dardanus venosus (Milne-Edwards) Yes Yes
Atrina rigida Lightfoot No No Emerita talpoida Say Yes Yes
Chione cancellata (Linne) No No Gecarcinus lateralis (Freminville) Yes Yes
Codakia orbicularis (Linne) No No Grapsus grapsus (Linné) Yes Yes
Donax variabilis Philippi Yes Yes Libinia erinacea (Milne-Edwards) Yes Yes
Gastropods: Macrocoeloma sp. Yes Yes
Busycon contrarium (Conrad) No No Menippe merceneria (Say) Yes Yes
Conus spurius atlanticus Clench No No Mithrax hispidus (Herbst) Yes Yes
Fasciolaria tulipa (Linne) Yes No Ocypode quadrata (Fabricus) Yes Yes
Littorina deh No No Pachygrapsus transversus (Gibbes) Yes Yes
Nassarius vibex (Say) No No Panopeus herbstii Milne-Edwards Yes Yes
Nerita spp. ae No No Petrochirus diogenes Linné Yes Yes
Pleuroploca gigantica (Kiener) No No Portunus spp. Yes Yes
Prunum apicinum Menke No No Sesarma cinereum (Bosc) Yes Yes
Pyrula sp. No No Stenorhynchus seticornis (Herbst) Yes Yes
Strombus alatus Gmelin No No Uca pugilator (Bosc) Yes Yes
S. gigas Linne Yes No Echinoderms
Cephalopods: Echinaster sentis (Say) No No
Octopus briareus Yes Yes Echinometra sp. No No
0. br (aredsteoo= No Yes Holothuria floridana (Pourtales) No No
O. joubini Robson Yes Yes Linckia guildingii Gray No No
Merostome Lytechinus variegatus (Leske) No No
Limulus polyphemus (Linné) Yes No Ophiothrix sp. ; No No
Oreaster reticulatus (Linne) No No
Crustaceans Fishes
Shrimps, mysids, lobsters: Acanthostracion quadricornis (Linné) Yes Yes
Alphaeus formosus (Gibbes) Yes Yes Adinia xenica (Jordan and Gilbert) Yes Yes
Gonodactylus sp. Yes Yes Cynoscion arenarius Ginsburg Yes Yes
Hyppolyte sp. Yes Yes Cyprinodon variegatus Lacepede Yes Yes
Mysidopsis almyra (Bowman) Yes Yes Fundulus similis (Baird & Girard) Yes Yes
Palaemonetes pugio (Holthius) Yes Yes Hippocampus erectus Perry Yes Yes
Panulirus argus (Latreille) Yes Yes Menidia beryllina (Cope) Yes Yes
Penaeus aztecus (Ives) Yes Yes Micropogon undulatus (Linnaeus) Yes Yes
P duorarum Burkenroad Yes Yes Monacanthus ciliatus (Mitchill) No No
Squilla empusa Say Yes Yes Opsanus beta (Goode and Bean) Yes Yes
Synalpheus brevicarpus (Merrick) Yes Yes Pogonias cromis (Linné) Yes Yes
Tozeuma carolinense Kingsley Yes Yes Sciaenops ocellata (Linnaeus) Yes Yes
Scorpaena brasiliensis Cuvier Yes Yes
Trachinotus caro (Linné) Yes Yes

and 4.8% from the exponential curve-fitting equations of the
first 20 weeks in figure 11. In all cases of weight and length
measurements, growth began to decrease in the period be-
tween 16 and 20 weeks, irrespective of temperature changes.
The length-weight relationship was calculated from the 1983
data and is shown in figure 13; Aronson (1982) gave com-
parable data for field-caught Octopus briareus.

In summary, Octopus briareus growth rivals that of any
fast-growing cephalopod (Forsythe and Van Heukelem, 1987)

and the data here are as comprehensive as for any species
(Forsythe, 1984). Previous studies indicate that temperature,
prey density and sexual maturation affect feeding and growth
(Boyle, 1987). Borer (1971) was able to correlate feeding in O.
briareus with prey density and temperature under fairly natural
temperature fluctuations. Studies in the open system in Miami
indicated that slowing of growth could also have been cor-
related with a drop in winter temperature late in the year and
the onset of sexual maturation. It is noteworthy that the 1983

26 AMER. MALAC. BULL. 7(1) (1989)

(6) Exponential
50

ML = .677e°'S"
(r2 = 9887)

ML = .0206t'*8”
(r2 = .9850)

Mantle length (mm)

10 20 30 40
Age (weeks)

Exponential Logarithmic

©

MW = 5.10e°'>*

= (r2 = .9957)
E
a
ao]
s
= Mw = .0395t'4””
6 (r2 = 9870)
=
10 20 30 40

Age (weeks)

Exponential Logarithmic

300 xp ~ 800 g
+ 700

TL= .1361t'°?®

TL = 24.07e°'%™
ane (r2 = .9690)

(r2 = .9896)

180

Total length (mm)

60

Age (weeks)

growth data are from eggs obtained in Eleuthera, Bahamas
where Aronson (1985) followed field growth of that same year
class. Our laboratory reared animals grew to very large size
and indicate that the small adult size of the individuals in the
Eleuthera population is not limited genetically but ecologically.

ALLOMETRIC GROWTH

Typically, the slope (b) or power exponent of the length-
weight relationship for animals lies between 2.5 and 4.0 (Brody,
1945; Brown, 1957). When b = 3.0, weight (or volume) is con-
sidered to be increasing as the cube of length (or linear size)
and is indicative of isometric body growth (Gould, 1966; Ricker,

®

Exponential Logarithmic
200 +
= AL, = 12.85e°'8"
e 160 + (72. = 9690)
= At
m 120+
Cc
= i
E
= 80 +
B t
ic 40+ AL, = .0058t? °°
I [19° (r2 = 9710)
ee rere area rere rere er rarer rare res rere
10 20 30 40
Age (weeks)

Exponential

AL, = 17.16e°'8
+ (r2 = 9727)

Logarithmic

AL, = .0197t' 87'°
+ 100 (r2 = 9760)

aR EAL A A SL IL Lon kee ekonomi
10 20 30 40

Age (weeks)

Figs. 6-10. Length versus age with mean, range and standard devia-
tion shown for each measurement period (1983 data). Compare with
figure 11. NOTE: The lines connect the mean values and are not
generated from the equations.

1979). When b > 3.0, weight is increasing at a rate greater
than that required to maintain constant body proportions, thus
indicating allometric body growth (Ricker, 1979). By these
definitions, the slopes of the equations in figure 13 indicate
that body growth of Octopus briareus is allometric throughout
half the life cycle.

To determine more precisely if growth was occurring
allometrically relative to any two of the body dimensions, the
length measurements were fitted by a regression to the
allometric equation Lz = aL,°. L, (the independent variable)
and Lz (the dependent variable) represent the two body
lengths being analyzed, ais a constant and b is the constant
of allometry. Both a and b are derived from the regression
(Simpson et al., 1960; Ricker, 1979). When b = 1, growth is
isometric with respect to the two lengths being compared,
meaning they are growing proportionally. When b > 1, the
Lz dimension is growing faster than the L; dimension, and
when 0 < b < 1 the converse is true. In either situation growth
is allometric.

The constant of allometry for each comparison is listed

—
Kp

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 27

Table 2. Growth in wet weight of Octopus briareus cultured through the life cycle in closed system aquaria in 1983. Growth rates are given
as instantaneous (inst.) and those calculated from measurement to measurement and overall (ovrl.) from day 14 to each measurement. Dou-
bling time (DBL) is the number of days needed to double in weight at the corresponding instantaneous growth rate.

Growth Rate

Mean Wet Range %/day g/day
Day No Weight S.D. DBL
(g) min. max. inst. ovrl. inst. ovrl. (days)
14 20 0.20 0.04 0.16 0.27 ---- ---- ---- = a---
28 19 0.50 0.07 0.39 0.65 6.43 6.43 0.03 0.03 10.78
42 19 0.98 0.16 0.68 1.32 480 5.61 0.05 0.05 14.45
56 18 1.75 0.37 1.09 2.39 4.16 5.13 0.07 0.09 16.65
70 15 3.49 0.63 2.29 5.04 4.09 5.07 0.17 0.18 14.13
84 15 8.07 1.67 5.06 11.40 6.00 5.26 0.48 0.42 11.56
98 15 14.03 3.39 9.34 22.69 3.95 5.04 0.55 0.71 17.56
112 14 25.02 5.41 17.19 36.63 4.14 4.91 1.03 1.23 16.76
126 14 45.33 11.74 30.26 72.93 4.24 483 1.92 2.19 16.34
140 14 68.49 19.91 40.60 113.04 2.95 462 2.02 3.16 23.51
154 14 121.78 40.28 70.43 210.34 4.11 457 5.01 5.56 16.86
168 14 183.71 48.19 117.78 296.09 2.94 442 5.40 8.12 23.60
182 14 281.77 71.15 175.00 438.20 3.06 431 8.61 12.13 22.69
196 13 362.50 70.53 267.60 519.10 1.80 4.11 652 14.91 38.52
211 11 450.48 98.58 335.20 698.10 1.45 3.91 6.53 17.61 4785
224 10 546.32 108.05 414.90 805.60 1.48 3.76 8.11 20.54 46.72
238 10 645.46 128.44 492.70 971.90 1.19 3.60 769 23.23 58.19
252 9 707.42 145.54 528.50 1054.70 0.65 3.43 463 24.24 105.87
266 6 746.70 48.15 665.70 804.10 0.39 3.26 2.88 24.32 179.59
281 5 826.54 69.64 726.20 901.90 0.68 3.14 5.60 25.72 102.35
295 5 877.28 82.29 775.20 963.20 0.43 2.98 3.73 26.13 162.88
309 7 811.86 183.86 510.00 984.00 -0.55 2.81 -4.49 2282 -125.21
324 7 869.50 239.29 432.42 1083.14 0.46 2.70 3.98 23.45 151.58
336 5 847.24 182.40 538.50 994.30 -0.22 2.59 -1.83 21.93 -320.72

in Table 3. The arms grow faster than other body parts dur-
ing the exponential growth phase up to day 126. Relative to
total length, the mantle length and width are growing at slower
rates (b < 1.0), while the arm lengths (arm pairs 1 and 3) are
growing isometrically with total length (b = 1.0). Consequently,
arm lengths are growing faster than mantle length (b > 1.0)
and are the major contributors to total length increases (Fig.
14). During the logarithmic phase (days 140 - 224), mantle
length and width grow faster than total length (b > 1.0), which
is partly a reflection of gonad maturation.

Therefore, the characteristic long arms of Octopus
briareus (Pickford, 1945) are a result of positive allometric
growth during the first 18 weeks of the life cycle. By com-
parison, Forsythe (1984) reported that O. joubini Robson
(which is morphometrically almost identical at hatching with
O. briareus) showed much less dramatic changes in body pro-
portions during growth. The proportion of arm length to total
length in O. joubini changed from 62% at hatching to 72%
at maturity, whereas the same proportions in O. briareus
changed from 50% to 78% (Fig. 14).

MORTALITY RATE IN CULTURE

Octopus briareus can be reared in individual containers
throughout its entire life cycle; survival is 70 - 90% throughout
the life cycle, the octopuses grow extremely fast and many
of them mate normally and can lay eggs for second genera-

Table 3. Constants of allometry (b) for both growth phases calculated
from allometric equations for total length (TL) relative to mantle length
(ML), mantle width (MW), first arm length (AL;) and third (AL3) arm
lengths (n=8).

Exponential phase Logarithmic phase

Comparison 14-126 Days 140-224 Days
b b
TL/ML 0.8405 1.1470
TL/MW 0.8764 1.1830
TL/AL, 0.9909 0.9083
TLIAL3 1.0390 0.9431
ML/AL, 1.1300 1.2620
ML/AL3 1.2030 1.0530

tion. Survival drops considerably when octopuses are mass-
cultured. There is usually a slow but steady mortality through-
out the life cycle (Fig. 15), although most mortality occurs
before a weight of 10 gm. Forty to 60% mortality by the adult
stage is common under these conditions. Causes of death
vary widely by experiment, but can include premature or non-
viable hatchlings, escapes, aggression, cannibalism, disease,
senescence and laboratory accidents. Escapes, aggression
and cannibalism are more common in O. briareus than in other
large-egged octopus species that have been reared in the
laboratory (Hanlon and Forsythe, 1985).

28 AMER. MALAC. BULL. 7(1) (1989)

(1) Exponential Growth Phase

-©

O- ---01983
16
| O----O 1983 , Caan
“oO aon
14 a J 4 &---2\1975
aaah 4 Vi
12] &--~4 1975 / Bel
— 4 = 4
oO 0 0481 7 >
1.0 W= 0.12166 4 ‘ 0
E ] (2 = 0.9968) o% / eee
@ So Cd / o a|
= 08 4 Pi p 401
x= .
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= W = 0.096009 046 Pa J. o
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=) 08 (r2 = 0.9962) “ Pa = © 30 |
so Pig = 4
/7 W= 0.065e0 0481 § 204
A (r2 = 0.9700) ° 4
o* 404
T ae 7 ie r T T: .-- Tr ™m rT a vr —
6 7 8 4 8 12 16 20 24 28 32 36 40 44 48
Age (weeks) Age (weeks)
Exponential Growth Phase cet
40.0 = 500
O---O 1983 a 4 r
35 0| 0-0 1981 / / [ WW = .00301 ML?°7
5 - a
a y / r= 9845 ~~
30.04 / / 100
= ! ;
E 9501 W = 0.121669 g /
2 (r2 = 9968) / /
2 = /
ra ; /
E 2004 iy =
o , = WW = 000313 ML3"3
= 1504 W=0.096e0 0461 re mA ms = 9947
(r2 = 0.9962) 7 , @ 10 es <
10.0 We So :
W = 0.065e0 0481 2
5.0 (r2 = 0.9700)
nates ee | T as 7 T
8 9 10 11 12 13 14 15 16 17 18 19 20 1.0
Age (weeks)
05
Logarithmic Growth Phase
900.01 o.-.0 1983 1 5 oe (4
800.0 7 OO 1981 / A 44 1 taiiit on eee
J 9 / 5 10 50 100
4000, 2 1978 lie F
/ / Mantle length (mm)
70 /
q 800.09 W= (1278 x 10-8)t410 Pe
E en (r2= 0.9800) A)’ W= (3.59 x 10°73. 76 DAYS OLD
© ; 2=
2 we 5 | |"? = 0.9861) 5 14 a Say
£ 4000 Y oo
> Ov $e
= 30004 Pes
ov:
200.0 4 “ W = (2.25 x 10°7)t3.88

(r2 = 0.9994)

T T T si | if
20 24 28 32 36 40 44 48
Age (weeks)
50% 61% 69% 78%

Fig. 11. Growth in wet weight versus time. The growth curves were derived from the given equations using the plotted data points, which repre-
sent mean weights. Age in days is t and the coefficient of correlation is r2. See Table 2 for details and compare 1983 data with figures 6-10.
Figures 11 and 12 reprinted from Hanlon (1983) and with permission of Academic Press with addition of 1983 data. Fig. 12. Growth rates ex-
pressed as percent increase in body wet weight per day. Growth rates were determined from the equation:
es log e Y2- loge Y,; X 100
to- t

where G is the instantaneous relative growth rate, Y; is the initial wet weight, Y2 is the final wet weight, t; is the age in days at Y,, and tz
is the age in days at Y2. All calculations are from the original data in figure 11. Each growth rate represents only growth between measurement
days (usually every two weeks) (Figs. 11 and 12 reprinted by permission of Academic Press Inc.). Fig. 13. Length-weight relationship calculated
from the 1983 data, calculated separately for each growth regime. Fig. 14. Allometric growth. During the exponential growth phase the arms
grow very fast relative to the rest of the body.

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 29

2G
OC,| 1 JOC,
. !
°C ne
8.0" -"-
H
P 7.6 Oc,
OC, + OC) oc] 1 /OC,
8.0 fie : ° ares ' ° aoe sae i ae
pH ; a = ae ahs
mg/l
mg/|
op)
LW
7)
=
oO wi
O>
ss Zz 60
oe)
fe)
> 20
@) 50 100 150

200
150
100 mg/l
50

200

150

100 mg/l
50

250 300 350 400

DAYS

Fig. 15. Typical results of 1983 large-scale culture experiment in which the octopuses were reared in large groups. Large numbers of animals
were taken out purposely on days 70 and 80. Note the high levels of ammonia, nitrite and nitrate in the latter portion of the experiment, in-
dicating that the animals are much more hardy than previously thought. OC ; 2 3 are different tank systems.

CANNIBALISM AND PATHOLOGY

Cannibalism is a common trait of Octopus briareus and
even in the very young stages the animals will completely can-
nibalize conspecifics when food is in short supply. This is not
merely an artifact of laboratory conditions; Aronson (1989)
noted six observations of cannibalism in his field study.

Mesozoan parasites occur in the kidneys of Octopus
briareus but do not cause mortality (Short, 1961). Fatal skin
ulcers (Fig. 16) were caused by Vibrio spp. that occur naturally
in seawater; full details of this disease can be found in Hanlon
et al. (1984). The initial cause of skin damage was the effect
of sucker marks when young animals aggregated during the

young stages of group culture. Octopuses grown in individual
containers in the same seawater system were free of disease.
The anti-bacterial compound nifurpirinol (Furanace®) was ef-
fective in stopping the progression of the ulcers and several
animals healed completely two months after treatment. Sucker
scars on the mantle of laboratory reared adults and octopuses
in nature do not seem to get infected similarly, indicating that
larger animals could be less susceptible to secondary
infection.

SENESCENCE
Senescence occurred in 10 - 12 months in all laboratory

30 AMER. MALAC. BULL. 7(1) (1989)

é

Fig. 16. One-month-old juvenile with severe skin ulceration of the mantle (whitish areas). The arms are normal. Fig. 17. Senescent adult oc-

topus. Note the poor condition of the skin of the mantle.

studies, although two animals lived particularly long: one
male lived 500 days in open-system culture and another male
lived 492 days in closed-system culture. Both animals attained
only a moderate size, and both had almost certainly mated
in the time period of 190 to 200 days. In the vast majority of
natural deaths males and females underwent a two to four week
period of deterioration, during which feeding was sporadic
and the skin, arms and internal organs degenerated (Fig. 17).
In most males, this deterioration occurred at varying periods
after mating and growth to a large size. In females it occurred
after egg laying and brooding. The most obvious manifesta-
tion of senescence was that the skin tone degenerated and
the skin became gray and-the papillae were inoperable. The
mechanisms of death are unknown but are probably linked
to the hormone system that regulates sexual maturation (Van
Heukelem, 1979; O’Dor and Wells, 1987). Rearing studies from
different brood stocks in different years show consistently that
the life span ranges from ten to 17 months; field data indicate
the same (Hanlon, 1983). Efforts to increase longevity by
feeding brooding females or rearing octopuses at constantly
warm temperatures with high food availability result in the
same mortality as brooding females without food or octopuses
reared in open systems with normal temperature fluctuations.

BEHAVIOR

The importance of behavior in describing and
understanding all activities of octopuses cannot be over
stressed. Octopuses are generally solitary until they mate.
Their soft bodies require that they avoid predator detection
during their daily foraging for food. They accomplish this by
camouflage, by operating mainly in the dark, by taking ad-
vantage of bottom relief for protection and, at last resort, by
threatening predators with specific body patterns or by eject-
ing ink and escaping.

Octopus behavior is complex. The central nervous

system (CNS) is large and organized into many discrete lobes
(Young, 1971). Their vision is superb (Messenger, 1981) and
they have demonstrated abilities of learning and memory
(Wells, 1978). In Octopus briareus these attributes are used
to compete in the high-density habitats associated with Carib-
bean coral reefs (Hanlon, unpub. data). Predators on oc-
topuses are fishes and mammals (e.g. Randall, 1967; Packard,
1972).

We attempt in this section to describe and explain the
main facets of behavior of Octopus briareus. The data are
based mainly upon laboratory observations (over 1400 hours)
but have been corroborated with field observations throughout
the Caribbean Sea (Hanlon, unpub. data). Laboratory obser-
vations were made mostly (> 60%) during the day, with
numerous observations made at night with the lights on, and
a few at night with a 30-watt red light. Distinctive body pat-
terns and behaviors of O. briareus that are useful in species
identification have been compared by Hanlon (1988).

MORPHOLOGY OF BODY PATTERNS

Anyone who has observed a live octopus takes im-
mediate notice of the changing color and texture of the skin
as well as the soft, supple body that can assume a variety
of shapes. The appearance of an octopus at any given mo-
ment is known as a body pattern, and its expression is
mediated by the remarkably well-developed eyes, CNS and
skin. The octopus is almost certainly color blind (Messenger,
1981), but it can blend with its background by adjusting the
expansion of its numerous, neurally controlled chromatophore
organs in the dermis of the skin. The chromatophore system
is regulated primarily by visual input to the eye. Together, the
eyes and skin constitute a system for camouflage that
matches luminance (Messenger, 1979). Below the
chromatophores in the dermis is a system of broad-band
reflecting cells (i.e. iridophores, reflector cells and

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 31

Fig. 18. Close-up photograph of the skin of an adult. Marked area indicates ‘‘visual unit’’ (scale = 1 mm). Fig. 19. Skin close-up of visual
unit (scale = 0.5 mm). Figs. 20, 21. Adult octopus sitting on a coral head. Several examples of ‘‘visual units’’ are circled in ink. Note various
component numbers. Fig. 22. Adult octopus in the Acute Mottle pattern. Note numbered components. Fig. 23. Large adult performing the
Parachute Attack maneuver as a speculative pounce on a small coral at night off Roatan, Honduras.

32 AMER. MALAC. BULL. 7(1) (1989)

leucophores) that reflects and scatters incident light of all
wavelengths. Taken together, the chromatophores and reflec-
ting cells are capable of producing a wide spectrum of visi-
ble light from the skin.

All known aspects of behavior are associated with
specific body patterns. Thus, one cannot adequately describe
behavior in cephalopods without describing the body patterns,
many of which are species-, sex-, age- and behavior-specific.
Packard and his collaborators (e.g. Packard and Sanders,
1969, 1971; Packard and Hochberg, 1977) developed a hier-
archical classification in which elements (e.g. chromatophores)
are grouped into units or skin patches, groups of units make
up specific components, different components form patterns
on the whole animal, and patterns are reflections of the whole
behavior of the animal. Packard and Hochberg (1977)
developed four general principles of patterning in
cephalopods, and the interested reader should consult that
paper for details. This classification of patterning has been
used to describe patterning in several cephalopods including
Octopus vulgaris (ibid.), O. burryi Voss (Hanlon and Hixon,
1980), Eledone cirrhosa (Lamarck) (Boyle and Dubas, 1981),
the teuthoid squid Loligo plei (Blainville) (Hanlon, 1982)
and the sepioid cuttlefish Sepia officinalis Linné (Hanlon and
Messenger, 1988). For standardization, capitalization is used

for the components (first letter only) and body patterns (first
letter of each word).

ELEMENTS: These are the smallest visible entities in the skin
that produce color or texture. In Octopus briareus, this includes
chromatophores (three color classes: yellow-orange, red-
brown, brown-black), reflecting cells and papillae. The
chromatophores (Figs. 18, 19) are small (approximately 0.011
mm retracted, 0.10 mm expanded) and dense in the skin (ap-
proximately 400 per mm? in adults). There are also extra-
tegumental chromatophores on the dorsal viscera of hatch-
lings; these expand and retract for several weeks posthatching
and are a conspicuous element of patterning in young oc-
topuses when the mantle is still translucent. The variety of
reflecting cells (i.e. iridophores and leucophores) has not been
studied in detail (i.e. with light and electron microscopy), but
one type produces the blue-green coloration that is a
distinguishing character of this octopus species. Papillae can
be produced all over the body; a generalization is that there
are short (1 mm) round (0.5 mm diameter) papillae, and long
(3 mm) round (3 mm diameter) papillae.

UNITS: These are difficult to define in Octopus briareus.
Packard and Hochberg (1977) originally depicted units as the

Table 4. Body patterns and their components in Octopus briareus (the numbered components are listed on most figures).

CHROMATIC COMPONENTS

Light

(1) Pupil margin
(2) White iris
(3) Iris margin

) Head bar

) Transverse mantle bar

) White patches

) White papillae arms
) White transverse arm bands

(4
(5
(6
(7
(8

head & mantle

Dark
(9) Pupil
(10) Dark iris
(11) Dark eye ring
(12) Reflective eyeball
(13) Branchial hearts
(14) Extrategumental chromatophores
(15) Dark hood
\ (16) Mottle
(17) Transverse arm bands
(18) Dark edge suckers

arms

TEXTURAL COMPONENTS
(19) Smooth skin
0) Coarse skin
1) Papillate skin

(2
(2
(22) Prominent mantle papillae

POSTURAL COMPONENTS
(23) Standing

(24) Protective posture

(25) Outstretched arms

(26) Interbrachial web spread
(27) Tucked in, curled arms
(28) Coiled arms

(29) Flattened head

(30) Raised head

(31) Distended mantle

LOCOMOTOR COMPONENTS
(32) Head bobbing

(33) Leaning

(34) Water jetting

(35) Inking

BODY PATTERNS
Chronic patterns (hours or days)
1. Uniform Light Phase
2. Uniform Light Blue-green Phase
3. Chronic General Mottle

5. Acute Mottle
6. Deimatic
7. Passing Cloud

Uniform Darkening

MANEUVERS

Acute patterns (Seconds or minutes)

Parachute Attack

Pincer Feeding Approach
Side Arm Attack
Countershaded Swimming
Copulation

Cleaning Maneuver

PaPrond >

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 33

static morphological array of elements in the skin (especially
the chromatophores). In O. vulgaris there is a conspicuous
morphological unit - a system of grooves that create obvious
skin patches, or ‘‘chromatic units,’ but this arrangement is
not seen in all octopuses, including O. briareus. The concept
of a ‘‘physiological unit’’ can also be considered (Packard,
1982), based upon neural control of the units by motor axons
originating in the CNS. In our study we limit our analysis to
small circular ‘‘visual units’ that are mainly physiological en-
tities generally appearing dark or light in various components
(Figs. 18-23). Each visual unit has a papilla in the center and
varies in size, but there are three basic size categories: 0.5
mm diameter with approximately 80 chromatophores; 1.5 mm
with approximately 700 chromatophores; and 3.0 mm with ap-
proximately 1500 chromatophores. Each visual unit also com-
prises an unknown number and arrangement of reflecting
cells such as the leucophores that reflect the bright white seen
in many components. We do not promote use of the term
“visual unit’ until detailed work is undertaken.

COMPONENTS: These are the recognizable and repeatable
parts that constitute the whole body patterns. Thirty-five are
listed in table 4 under four categories: (1) chromatic, (2) tex-
tural, (3) postural and (4) locomotor. As explained by Packard
and Sanders (1971), components may be expressed in a wide
variety of combinations. Some components commonly go
together while others are mutally exclusive. Collectively they
confer upon the animal the ability to show a highly diversified
range of body patterns.

Chromatic components are those concerned strictly
with color. They are conspicuous and well defined and occur
repeatedly in the same relative position on the body. They
are recognizable because of contrasting light and dark areas.
The light components result when the overlying chromato-
phores are retracted and light is reflected from the underly-
ing leucophores or iridophores. The dark components result
from light that is reflected from and transmitted through the
pigment granules of expanded chromatophores. Chromatic
components are physiological entities that reflect selected
neural activity because individual chromatophores are con-
trolled directly from the CNS (cf. Messenger and Miyan,
1986).

The components are numbered (Table 4) and most are
self-explanatory and indicated on the figures. Figure 24
depicts the arrangement of seven chromatic components that
are associated with the eye. The appearance of the eye fluc-
tuates constantly, and it can appear prominent or obliterated
depending upon the combinations of these components that
are expressed at any given moment. The iris can be either
light (Comp. 2) or dark (Comp. 10) depending upon the degree
of expansion of the iris chromatophores. The pupil of the eye
always appears black. Depending upon the quantity of inci-
dent light, the pupil can appear as a thin horizontal slit or a
circle. The outer perimeter of the eye is generally the same
color as the head, and with expanded chromatophores it forms
the Dark eye ring. The region above the eye can also be
papillate (Comp. 21). In young animals, the presence or
absence of expanded chromatphores on the outer eye ring

Fig. 24. The chromatic components of the eye. Note the range of
expression. Not all pupils were printed horizontal. A, Dark eye ring;
B, Pupil margin; C, Pupil; D, Iris; E, Iris margin.

determines how obvious the Reflective eyeball will appear in
hatchlings. The Dark eye ring can make the eye appear larger
by matching the color of the eye or contrasting with the Iris
margin. The Pupil margin and Iris margin are thin rings that
enhance the contrast of the pupil or eye. Some representative
illustrations of components common in young O. briareus are
shown in Figs. 25-32.

The White patches (Figs. 20-23; Comp. 6) are irregular-
ly shaped and made up of several circular visual units in which
the chromatophores are retracted. The white Head bar (Figs.
26, 29, 30; Comp. 4) consists of an irregular, transverse row
of white patches between the eyes. The Transverse mantle
bar (Figs. 29, 30, 33; Comp. 5) is irregular in shape and con-
sists of a series of white patches each with a White papilla
(Comp. 7). White transverse arm bands (Figs. 20-23; Comp.
8) are fairly regularly spaced and are made up of groups of
White patches.

The dark components Branchial hearts (Figs. 27, 29;

34 AMER. MALAC. BULL. 7(1) (1989)

Fig. 25. Twenty-four-day-old juvenile. Note the extrategumental chromatophores (14) of the arms (two rows), head and visceral mantle. Fig.
26. Thirty-four-day-old juvenile. Note the newly developed Head bar (4). Fig. 27. Thirty-eight-day-old juvenile. Note newly developed iridophore
splotches (arrow), Reflective eyeballs (12) and Branchial hearts (13). Fig. 28. Thirty-day-old young showing unilateral expression of
chromatophores on the mantle. Fig. 29. Fifty-day-old juvenile in Uniform Light Phase. Note the formation of the Transverse mantle bar (5)
and the Head bar (4) by aggregations of iridophore or leucophore splotches. Fig. 30. Two young octopuses, the one in the foreground showing
Head bar (4) and Transverse mantle bar (5). Fig. 31. Young octopus on a reef showing Outstretched arms (25) and Raised head (80).
Fig. 32. Young octopus in Uniform Darkening pattern showing the locomotor component Leaning (33) while it sights a prey organism.

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 35

Comp. 13) and Extrategumental chromatophores (Figs. 4, 25;
Comp. 14) are evident only several weeks posthatching when
the mantle is translucent. Dark hood (Fig. 34; Comp. 15), in
its fullest form, includes all of the head, eyes and mantle, but
can only cover the head and the area in front of the eyes.
The dark Mottle (Figs. 20-23; Comp. 16) can be expressed
as large circular patches with all chromatophores expanded
or as irregular reticulations. The dark Transverse arm bands
(Figs. 22, 23; Comp. 17) are irregularly shaped and often not
well developed. The bands extend onto the web in the form
of parallel dark streaks. Dark edged suckers (Fig. 35; Comp.
18) enhance the white suckers.

Most of the remainder of the components are self-
explanatory or evident in the figures. The Standing posture
(Comp. 23) is shown in figure 36. Outstretched arms (Comp.
25) are seen in figures 31 and 37. Tucked in, curled arms (Figs.
35, 38; Comp. 27) protect the delicate arm tips. Prominent
mantle papillae (Comp. 22) are about 3 mm high (Fig. 33) and
their placement is illustrated in figure 39. The Protective
posture (Fig. 40; Comp. 24) is a defensive posture in which
the suckers and sometimes the mouth (i.e. the animal’s
weapons) face an intruder, thus protecting the vulnerable head
and mantle. Females brood eggs in this posture (Fig. 5).
Coiled arms (Fig. 41; Comp. 28) maximize the web spread.
Various postures are assumed in swimming and these are
illustrated in figures 42-44. In Distended mantle (Fig. 45;
Comp. 31) the mantle is full of water and the animals holds
its breath. Vertical head bobbing (Comp. 32) is used during
prey fixation. A typical posture with Coiled arms (Comp. 28)
is shown in figure 46. The unusual Flattened head posture
(Comp. 29) is illustrated in figure 47. The use of these and
the other components in body patterning will be explained
in the following sections.

BODY PATTERNS AND MANEUVERS: Table 4 lists those
observed in Octopus briareus. We have listed seven basic pat-
terns under two broad categories: chronic and acute, depend-
ing upon their duration. Chronic patterns are used for con-
cealment, while acute patterns are used in inter- and in-
traspecific encounters while the octopus is out of its lair and
moving on the substratum.

Uniform Light Phase (Figs. 27, 29, 35) is a chronic pat-
tern observed frequently. It is characterized by no dark com-
ponents, leaving a uniform background of white, dull yellow
or brown; usually there are uniformly raised papillae. Uniform
Light Blue-green Phase is seen in the field over light sandy
patches around reefs. No chromatophores are expanded, and
the resulting light blue-green tint has a glowing effect that is
a result of reflection from the various reflecting cells. All
papillae are usually raised uniformly producing Coarse skin.
This same blue-green tint is present over much of the body
during the Deimatic pattern. Chronic General Mottle (Figs.
20-23, 31) is an extremely common pattern that is variable
in form. The head and mantle have more circular light and
dark patches while the arms are characterized mainly by
Transverse arm bands. In general, laboratory reared oc-
topuses showed less chromatic expression than field-caught
animals, and field-caught animals maintained for long periods

showed less-intense patterns over time.

Acute patterns are by definition short-lived and can be
generally regarded as immediate responses to stimuli (e.g.
predators, prey, conspecifics). Uniform Darkening (Fig. 37) is
characterized by the uniformiy maximal expansion of ali
chromatophores, resulting in an overall dark brown colora-
tion. The pattern results when the octopus is stressed, as
when approached closely by another octopus, a predator or
a human observer. The skin texture can be either smooth or
sculptured by varying degrees of raised papillae. Variations
include the presence of white eyes (roughly one-fourth of the
time) or some very dark mottle in the form of dark reticula-
tion. A more striking variation is a blue-green metallic sneen
that apparently is produced by the expression of iridophores
on raised papillae.

Acute Mottle (Fig. 22) is a variegated pattern that is
characterized by the components Mottle, White patches,
Papillate skin and Interbrachial web spread. It is often accom-
panied by dark eye components. The pattern is used when
the octopus is startled by a nearby object such as a large prey
organism, a predatory fish, another octopus or a human
observer. In some behavioral contexts, this pattern can be con-
sidered a precursor to the Deimatic pattern.

The Deimatic pattern (Fig. 41) is similar in form and
expression to that described for other octopods (first coined
as ‘‘Dymantic’’ by Young (1950) to mean warning or frighten-
ing display, but deimatic has the same root and is in wider
use). The octopus flattens its head (Comp. 29) and mantle
dorso-ventrally, the arms are tucked in and curled (Comp. 28)
and the interbrachial web is spread slightly (Comp. 26). Con-
currently the entire body surface turns pale white except for
the Dark eye ring, Dark iris and expanded Pupil. Partially
raised papillae form Coarse skin. This display is elicited only
by a very sudden, intense, close rush by a large object or
predator. The intensity of the pattern depends upon that of
the stimulus as well as the reaction of individual octopuses.
In situations of increasing intensity, the order of appearance
of the components is: (1) expanded Pupil, Dark iris and Dark
eye ring; (2) Flattened head; (3) Arms tucked in, curled; (4)
paling of arms and web; and (5) complete Deimatic.

Passing Cloud (Figs. 38, 46, 48) is a dynamic pattern
in which the interbrachial web is spread (Comp. 26) to its
fullest and the arms are coiled (Comp. 28) upwards to pre-
sent the greatest possible surface area. The body is held in
this posture while the octopus glides forward slowly.
Simultaneously, a unilateral chromatic effect occurs as alter-
nate clouds of dark brown and white, originating at one eye,
travel outward to the periphery of the mantle, web and arms.
New waves originate at the eye and side of mantle
simultaneously every second, and it takes about 1.5 secs for
each wave to reach the arm tips. This pattern is always shown
laterally towards another octopus and can last several minutes
and be repeated many times in succession.

Unilateral variations result when a different pattern is
present on each side of the body. Generally the side toward
a stimulus is dark while the side away is light. Newly hatched
octopuses also can do this (Fig. 28). This pattern resulted
when either a very large crab or another octopus was in the

36 AMER. MALAC. BULL. 7(1) (1989)

Fig. 33. Laboratory adult in a light brown Uniform Light Phase. Note two of the three Prominent mantle papillae (22). Fig. 34. Expression
of Dark hood (15) while sitting in the raised head posture with Smooth skin (19). Fig. 35. Adult octopus in Uniform Light Phase. Note Dark
edged suckers (18) and Coarse skin (20). Fig. 36. The postural component Standing (23) in a sand/seagrass area off Eleuthera Island, Bahamas.
Fig. 37. The acute pattern Uniform Darkening and an example of Outstretched arms (25). Fig. 38. The Passing Cloud pattern, with a wave
of expanded chromatophores passing over the arms, which are held tucked in and curled (27).

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 37

same tank with an octopus (Fig. 48). The former instance was
observed 51 times; however, in two instances the opposite
reaction occurred - the dark side was away from the stimulus.
In one instance when a large crab was put in the tank, the
octopus showed the chromatic components of Deimatic
toward the crab, while showing Uniform Darkening on the
other side.

Six behavioral maneuvers are noteworthy. The
Parachute Attack (Figs. 49, 50) is associated with foraging and
feeding. Sinel (1906) first described this motor action pattern
in which the octopus ‘“‘rises above its victim, and with ten-
tacles so out-stretched that the web that joins them part of
their length forms a parachute, it descends like a cloud on
its victim.’ The general body pattern is Dark hood and white
arms with smooth skin. These chromatic and textural com-
ponents of the pattern appear just when the octopus has posi-
tioned itself above the prey and is beginning to descend upon
it. Upon descent, the arms and interbrachial web are spread
rapidly in parachute fashion, and the octopus settles on the
prey and entangles it; the octopuses then immediately went
to Uniform Darkening. Several variations were observed (Fig.
23; Hanlon, unpub. field data).

In the Pincer Feeding Approach (Fig. 45) the octopus
is in a brown Uniform Light Phase, often papillate (Comp. 21).
This maneuver is used to seize prey. The second pair of arms
curve forward during the approach, and the fourth pair ex-
tend forward from underneath. The mantle is distended
(Comp. 31) while the octopus holds its breath and moves
toward the prey. The prey is grabbed in one motion as the
pincer (arms 2) is closed and the fourth pair of arms shoots
forward.

Side Arm Attack is used when prey are close. The arms
on the side toward the prey coil back, with the suckers out-
ward. Three or four arms extend rapidly above the prey then
grasp it and pull it into the web. The body pattern is usually
Uniform Darkening with Coarse skin.

During countershaded swimming in a backward direc-
tion (Fig. 42) the dorsal body surface is in brown Uniform Light
Phase while the ventral surface is pale. The skin texture is
coarse (Comp. 20) and iridescence is usually present from
the reflecting cells. This pattern provides the necessary com-
ponents for effective countershading of a swimming organism
(Cott, 1940).

Mating behavior and copulation (Figs. 51-55) were
observed and three postures were noted: (a) the male most
commonly sat atop the female with his arms and interbrachial
web covering the female’s mantle and head, and (b) occa-
sionally the female rotated around from her posture described
in (a) until the oral surfaces of her suckers were against the
oral surfaces of the male’s suckers, (c) the male and female
sat about 10 cm apart while the male extended his hec-
tocotylized arm toward the female. During copulation the
male’s hectocotylus was inserted into the female’s mantle cavi-
ty. Coloration and skin texture were variable. Brown Uniform
Light Phase and Chronic General Mottle were the commonest
patterns. During two matings the males turned pale white and
showed Dark hood (Comp. 15) for short periods. Iridescence
on the skin was common in all patterns.

Three Cleaning Maneuvers were observed. Commonly
an octopus would shed the sucker discs by twirling its arms
against the body and blowing them away with jets of water.
A second maneuver was performed by females after mating;
they would rapidly move the arms inside and on the outside
of the mantle. Finally, females cleaned eggs in their lair by
continually grooming the egg capsules with their arm tips.

ONTOGENY OF PATTERNING

Figure 56 gives the times of appearance of the com-
ponents, patterns and maneuvers of Octopus briareus cultured
in the laboratory. At hatching, there are no iridophores or
leucophores evident in the skin (they begin to appear at two

Fig. 39. Diagrammatic representation of: A - Head bar, B - Transverse
mantle bar, C - Prominent mantle papillae. Two prominent eye papillae
are also indicated. Fig. 40. Protective posture. Fig. 41. Deimatic pat-
tern. Fig. 42. Backwards swimming. Fig. 43. Backward medusoid
swimming. Fig. 44. Forward swimming. Fig. 45. Pincer Feeding Ap-
proach to a small crab. Fig. 46. The acute pattern Passing Cloud
being shown unilaterally on the right. Stippled areas indicate suc-
cessive waves of chromatophore expansion.

38 AMER. MALAC. BULL. 7(1) (1989)

Fig. 47. Chronic General Mottle in a laboratory reared adult reared in isolation. Note Flattened Head (29). Fig. 48. The Passing Cloud pattern
being shown unilaterally (on the animal’s right side) in an animal approximately 200 days old. Fig. 49. The Parachute Attack in a young oc-
topus (about 60 days old). Fig. 50. The Parachute Attack maneuver onto a crab in an animal 144 days old; note Dark hood (Comp. 15).

weeks) and the chromatophores are relatively sparse.
Therefore, patterning is limited for the first two months or so.
During this early period the Extrategumental chromatophores
are important, as are the Reflective eyeballs and Branchial
hearts.

Newly hatched Octopus briareus appear to be restricted
to four general body patterns. The first is the chronic pattern
Uniform Light Phase in which the skin is translucent white,
the eyes are prominent and silvery-blue, the visceral organs
appear pinkish through the mantle, and the dark Branchial
hearts show through the mantle and produce the effect of two
false eyespots at the posterior end of the mantle. After ap-
proximately four weeks the mantle becomes thicker and more
opaque, and Uniform Light Phase becomes more adult-like
in appearance because the internal organs are not obvious.
This pattern is common when young animals rest on a light
or white object and when they are swimming.

The second chronic pattern is characterized by the full
expansion of the Extrategumental chromatophores on the
arms and viscera (Figs. 4, 25) while most or all of the other
chromatophores on the body are retracted. This pattern was
most often seen when the animal was sitting on a dark ob-
ject or when the octopus was moderately excited upon sighting

a moving object nearby.

The third common pattern was Uniform Darkening in
which all chromatophores were expanded maximally. The
overall color was dark brown and the pattern rarely lasted
beyond two minutes. This pattern was observed when the oc-
topus was very excited, as when attacking a crab or when
startled by a nearby object.

The fourth pattern observed in animals younger than
three weeks was unilateral Uniform Darkening. The dark pat-
terning was always seen on the side of the octopus facing
another octopus or a larger crab that had been put inas a
prey organism.

Some gradual changes take place between days 20
and 50. The number and density of chromatophores begins
to increase and papillae develop at about day 30. The first
to appear are single large papillae above each eye. By day
30 some iridophore cells first appear on the mantle as small
(0.5 - 1.0 mm) isolated patches of reflective silvery-blue (Figs.
27-29). They then soon appear over the heads and arms, and
groups of them begin to form the chromatic components Head
bar, Transverse mantle bar and White transverse arm bands.
An example of a small animal just at this transition can be
seen in figure 29. A modified form of Passing Cloud has been

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 39

Fig. 51. Male (right) approaching a female just prior to mounting her. Fig. 52. Copulation. Male (right) mounting the mantle of the female.
Fig. 53. Copulation. Male (left) showing incomplete Dark hood (15), covering the female (arrow) with the interbrachial web.

40 AMER. MALAC. BULL. 7(1) (1989)

Fig. 54. Hectocotylus, third right arm modified for spermatophore transfer. Fig. 55. Copulation. Male is at the top and female at the bottom.
Arrow indicates the hectocotylus of the male inserted into the female’s mantle cavity.

seen as early as 30 days. The various components of the eye
generally begin to appear between days 40 and 60. The
Deimatic pattern is fully expressed at about day 100 and the
typical form of Passing Cloud was not observed until day 210.
However, it is likely that the animals are capable of express-
ing it before this time.

Copulation was not seen before six months of age. At
the end of the life cycle senescence sets in and the skin
begins to deteriorate. Most of the components and patterns
are affected during senescence, and the common body pat-
tern at this period is a variant of Uniform Light Phase.

LOCOMOTION AND EXPLORATORY BEHAVIOR

Octopus briareus moves by four principal methods:
crawling, backward swimming, backward medusoid swim-
ming and forward swimming (Figs. 42, 43, 44). Hatchlings are
capable of crawling and backward swimming. The animals
usually only swim when they are excited, and they do so in
the acute pattern Uniform Darkening and often squirt three
or four pseudomorphs of ink as they move backwards.

Medusoid swimming and forward swimming were only
observed later in the life cycle.

Exploratory behavior was common in octopuses of all
ages. When an octopus is placed in a new tank or an object
is placed in its home tank, the animal will usually first withdraw
into the Protective posture and then soon investigate new ob-
jects by extending one or several arms cautiously. The arms
can stretch a great distance and the animal is thus able to
use tactile and chemosensory organs in the suckers to ob-
tain information about new objects. Eventually the animal will
touch all objects in its tank and move around to investigate
them more carefully. Octopuses also use vision in exploratory
behavior. They will often lean in the direction of interest to
obtain better sight of an object before leaving their lair.

INTRASPECIFIC INTERACTIONS

Octopus briareus is a solitary animal for most of its life
cycle. During the first few weeks posthatching, the young
animals tolerate conspecifics and sometimes even aggregate
in group-culture conditions. However, they soon become in-

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 41

tolerant of conspecifics and cannibalism is common, especial-
ly during times of food shortage. When the gonads are ripe
the animals will readily mate, but they then separate and do
not form permanent mating pairs.

AGONISTIC BEHAVIOR: There is no evidence from laboratory
rearing that young octopuses are territorial or maintain a
permanent home. From hatching, octopuses seek shelter
such as empty shells, but Octopus briareus is not strictly noc-
turnal and can be seen moving about feeding both during the
day and night. Young animals have been observed feeding
on the same piece of shrimp meat. Intraspecific aggressive
behavior was first evident at five to six weeks of age. The first
interaction was observed at day 42 when two small octopuses
fought each other for five seconds. They both remained in
the Uniform Light Phase pattern with their arms folded
backward and interbrachial webs spread, and moved forward
bringing the buccal masses together. In a similar instance,
two octopuses of the same age approached each other in the
Uniform Darkening pattern and extended two arms each
towards each other. In some cases, an intruder was able to
remove an octopus from its den by grappling with it. Fighting
over food was common, especially during poor food availabili-
ty or when the animals were approaching two months of age.
For example, one young octopus was observed to pull the food
from the web of another and then return to its den. The oc-
topus that lost its food followed the first to the den and was
attacked, with the result that its fourth right arm was torn off.
Some form of hierarchy was apparent, and size and ag-
gressiveness were probably key elements in its structure.
By two months of age there were some examples of
a dominance hierarchy based upon size. The largest animals

appeared to have the primary choice for den selection and
feeding. This hierarchy remained constant when the animals
were moved to new surroundings.

Rearing conditions strongly affect the quality and quan-
tity of intraspecific interactions. If the animals are well spaced
and there is an excess of hiding places and food, then in-
teractions are not numerous or violent. Under more natural
conditions the majority of interactions between octopuses do
not end in fighting, but in displays and stylized attacks.

The acute pattern Passing Cloud appears to be used
in establishing dominance. If an octopus does not respond
to the Passing Cloud pattern, then the displaying octopus will
often touch the other. In many cases this results in the
subordinate animal fleeing or moving into the Protective
posture. In other cases, a bout can ensue in which the oc-
topuses entangle their arms attempting to maneuver on top
of one another. In this position the eventual winner will wrap
its arms and web around the mantle to restrict breathing.
Sometime the attacks are made in a side-arm fashion with
only two or three arms from each animal engaging in the bout.
In some cases, the subordinate animal will autotomize an arm
to facilitate rapid escape and the victor may eat the captured
arm. After intraspecific bouts, it is not uncommon for circular
gray wounds to be left on the mantle, presumably from the
effect of the suckers.

There was one documented instance in which the
dominant/subordinate relationship reversed over time. A male
and female had been reared in the same tank from hatching
to 100 days. Both had the circular scars on the body indicating
that bouts had taken place, but no dominance was observed
in either animal. Within several days, however, the male
became strongly dominant, feeding first and causing the

COMPONENTS
Chromatic
Light:
White pupil margin (1)
White iris (2)
White iris margin (3)
White head bar (4)
Transverse mantle bar (5)
White patches (6)
White papillae (7)
White transverse arm band (8)

1
\
\
|
\
|
|
\
\
{
\
\
|
Dark: H
Dark iris (10) t
Dark eye ring (11 |
Reflective eyeball (12) 6 —
Branchial hearts (13) ( o——----
Extrategumental chromatophores (14) |
Mottle (16) |
Transverse arm bands (17) |
|
!
|
\
|
|
|
\
\
\
I
|
|
|
{

Textural
Papillate skin (21)

Postural and Locomotor
Protective posture (24)
Interbrachial web spread cs)
Tucked in, curled arms (2
Head bobbing (32)
Inking (36)

BODY PATTERNS
Chronic

Uniform Light Phase (1)
Uniform Light Blue-green Phase (2)
Chronic General Mottle (3)

Acute |
Uniform Dark Phase (4)
Acute Mottle (5)
Deimatic (6)
Passing Cloud

Maneuvers
Parachute Attack (1)
Side Arm Attack (3) !
Copulation (5) '

—_ presence of the component or pattern in its full, normal expression
= presence in an altered or less-complete expression

Fig. 56. Development of patterning in Octopus briareus.

|
90 100 110 120 130 140 150 160 170 180 190 200 210 220 230 240 250 260 270 280

' | | | I | | | | ( | | i i | | | ( ( |

Number of days after hatching

42 AMER. MALAC. BULL. 7(1) (1989)

Table 5. Visual antipredatory adaptations of Octopus briareus (see Table 4 for pattern descriptions).

Adaptive coloration and behavior

Primary defense ¥ - concealment from predators
nocturnally active
remain motionless
general color resemblance”
countershading*
disruptive coloration”
concealment of shadow*

Body pattern

Purported effect

all patterns

all chronic patterns
all chronic patterns
Uniform Light

Chronic General Mottle

all chronic patterns

harder to see at night

predator is not attracted by motion
match substrate

blend with water column

obliterate body form

blend body outline with substrate

Secondary defense ¥ - make a predator hesitate
flash behavior*

flight*

Inking
Deimatic

Uniform Darkening
Passing Cloud
Acute Mottle
Protective posture
+ Water Jetting
Uniform Darkening
+Inking + Jetting

predator is startled and loses sight of prey

pattern, posture change and apparent size increase
bluff predator

pattern change confuses predator

rapid color change confuses predator

bold pattern change confuses predator

show the weapons, protect vital organs and
startle predator

predator is startled and loses sight of prey

Tertiary defense - misdirect a predator’s attack
deflective marks*
diversion behavior¥

Branchial hearts
Uniform Darkening
+ Inking + Jetting

misdirect predator’s attack with false eyespots
predator attacks ink blob, becomes disoriented
and loses sight of prey

terminology: *Cott, 1940

italics: components

WwEdmunds, 1974

female to relinquish captured crabs. However, during the next
30 days the female grew faster, became larger and subse-
quently became the dominant member of the pair. Other fac-
tors may have been involved such as a hormone change in
the female, whose gonads had enlarged during this time.

REPRODUCTIVE BEHAVIOR: Mating has been observed 12
times in the laboratory and once (Hanlon, 1983) in the field.
In all cases there was little or no courtship behavior, mating
appeared to be male dominated, and both males and females
would mate multiple times with different partners. Animals that
had been reared in isolation would readily mate when placed
in a tank with another mature octopus. The typical mating en-
counter was as follows. After being placed together in the
same tank, the male would advance across the aquarium (Fig.
51) and climb on top of the female, sitting on top of her man-
tle with his arms draped around the mantle and head (Fig.
52). The interbrachial web between the male’s first pair of arms
often covered the female’s eyes (Fig. 53). Most matings lasted
30 to 80 mins, but in two cases mating lasted 150 and 180
mins. During this time the male’s hectocotylized third right
arm (Fig. 54) would eventually be inserted into the mantle cavi-
ty of the female (Fig. 55) and a very long spermatophore
(sometimes longer than the mantle length) would be trans-
ferred to the oviduct of the female. Sometime during mating
it would not be unusual for long fragments of spermatophores
to be seen floating in the vicinity of the octopuses. The female
would occasionally struggle during mating, but termination
only occurred when the male released her and the two would
part quickly. Counts of ventilation rate of both partners were

of patterns

made to see if there were increases associated with transfer
of spermatophore to the oviducts (see Wodinsky, 1973 for Oc-
topus vulgaris). Five pairs were monitored and although oc-
casional increases in rate were observed (e.g. from 25 - 35
ventilations per min), there was no trend to indicate that ven-
tilation rate had anything to do with any specific aspect of
mating. At the termination of mating the females usually had
the distinctive sucker marks left on their mantle.

It is noteworthy that in most mating observations the
male was smaller than the female; therefore, some factor other
than size was important in domination of mating activity. In
the field observation of mating (Eleuthera Island, Bahamas)
a female 85 mm ML was mated by a male 53 mm ML during
mid-morning. This substantially smaller male completely
dominated the entire sequence - swimming across the bot-
tom, mounting the female and mating her for 58 mins. The
remarkable facet of this mating is that only minutes before
this was observed, the 85 mm ML female had been found
under a sponge eating the remains of another male that was
53 mm ML (Hanlon, 1983). Therefore, the receptivity of
females apparently can change within hours and could be
associated with their state of hunger, the degree of ag-
gressiveness of the male or perhaps some hormonal or
pheromone factor.

No single body pattern was associated with mating.
In several cases the male would be in a pattern in which the
arms and web were in Uniform Light Phase and the head and
mantle were uniformly dark brown. However, in most cases,
both animals were either in Uniform Light Phase or Chronic
General Mottle. In the field observation, the mating pair was

HANLON AND WOLTERDING: OCTOPUS BRIAREUS 43

initially in Uniform Darkening, but then gradually returned to
Uniform Light Phase in which they were a light brown color.

In all but one of the 13 mating observations the animals
did not seek cover or protection. Even in the field the oc-
topuses mated during the day in the open part of the reef.
This seems to be a very dangerous way to conduct an im-
portant part of the life cycle, but it remains to be proven
whether all animals in nature mate during the day in the open.
Mating with a larger female could be risky for a male Octopus
briareus. The danger of being captured and eaten if she is
non-receptive could be greater than the risk of being sighted
by a passing predator. Thus it can be to the male’s advan-
tage to mate during the day in the open where he can more
effectively monitor visually the receptivity of the female and
have room to escape if necessary.

In one laboratory observation, the male remained
within its den and extended his hectocotilized arm toward the
female sitting motionless on top of a rock about 10 cm away.
Copulation was observed for approximately 10 min, at which
time the female flashed Uniform Darkening and reached for
the male, who inked and fled.

In all observations of mating the animals were 200 to
250 days old, or in a size range of approximately 50 to 100
mm ML. This conforms generally to the time at which Octopus
briareus is thought to become mature. The hectocotylus has
been found in males as small as 27 mm ML. In laboratory
reared males the earliest observation of the appearance of
the hectocotylus was in four males between 40 and 50 mm
ML (133 days old, 26 - 59 g). Available evidence indicates that
females become mature at a similar size and age as males.
Examination of females in the University of Miami Museum
indicate that ripe ovaries are found in animals in the range
of 35 to 81 mm ML, and females 45 to 120 mm ML have laid
eggs. Conversely, females 18 to 45 mm mantle length are
generally immature (Hanlon, 1983).

Sperm storage can be as long as 100 days before egg
laying. Female behavior begins to change prior to egg lay-
ing: they reduce their food intake and they find or construct
a Suitably protected lair in which to lay the eggs. Females con-
stantly guard and clean the eggs and can only be separated
from them forcibly. The females appear not to forage for food,
but will often eat during egg brooding by grabbing crabs that
stray close to the lair.

Since males and females mate promiscuously, it is like-
ly in nature that individual females receive sperm from several
males. Sperm storage is thought to take place in the oviduct
or the oviducal gland, and it would be interesting and infor-
mative to know if there is any form of sperm competiton and
what effect it would have on the genetic makeup of the
population.

Unusual behavior by the female was noted after one
copulation. Five minutes after termination of mating the female
inhaled, raised her mantle straight up for five to eight seconds,
then exhaled forcibly while lowering the mantle to its normal
position. Two minutes later, while sitting in the head-high posi-
tion, she furiously curled her wriggling arms back and forth
over the mantle for 30 secs and then continued this behavior
every four minutes. In the interim she would place from one

to three arms into her mantle cavity for approximately 20 secs
then suddenly withdraw them with a contraction of the man-
tle. This behavior continued for seven hours and the ventila-
tion rate remained high, from 38 to 42/min. Thereafter, her
behavior was completely normal. This behavior remains
enigmatic.

Females that have been mated will frequently die
without laying eggs, despite producing a large number of eggs
in the ovary. In a normal female, eggs constitute one-third of
the mantle cavity, while in abnormal females the eggs con-
stitute roughly two-thirds of the mantle cavity. In the latter case
the internal organs are often compressed anteriorly into a
small volume. Several dissected specimens showed that the
proximal oviducal aperture was closed. Eggs pass via this
aperture from the ovary to the proximal oviduct. The result
is that eggs cannot be laid, and the eggs begin to decom-
pose in the ovary. Presumably it is some artifact of the
laboratory that results in this condition.

INTERSPECIFIC INTERACTIONS

FEEDING AND ATTACK BEHAVIOR: Three distinct feeding
maneuvers were observed in the laboratory: Parachute At-
tack, Side Arm Attack, Pincer Feeding Approach. The
Parachute Attack is illustrated in figures 23, 49, 50. Octopuses
as young as 22 days began to use Parachute Attack and by
70 days it was a commonly used attack maneuver. The young
animals would often miss the prey entirely by descending
short of the prey. Head bobbing (Comp. 32) was first record-
ed at this time. The animals would bob their heads before
the attack, presumably to aid in monocular paralax. A typical
sequence of feeding would be as follows. As the crab is
sighted, the octopus raises its head turning one eye toward
the prey; respiration rate increases and the eye and head
region or the entire body would darken. Head bobbing en-
sues as the octopus leans toward the prey. The arms would
coil beneath the body in preparation for the attack. The oc-
topus would then launch itself forward in the Parachute At-
tack sequence. Upon capturing the prey, the octopus goes
to Uniform Darkening for approximately ten seconds before
it reverts back to Acute Mottle and returns to its den to con-
sume the prey.

The Side Arm Attack sequence was used to seize prey
nearby. The closest three or four arms would be curled and
rolled back, then rapidly extended outward and upward, seiz-
ing the prey and pulling in into the web. Uniform Darkening
is associated with this maneuver. Newly hatched octopuses
use some parts of this attack sequence, but the fully
developed attack and its associated body pattern was not
observed until 44 days posthatching.

The Pincer Feeding Approach (Fig. 45) was observed
less often than the other two methods. The octopus faces the
prey and, with the second arms extended forward and the
first and third arms extended outward, would distend the man-
tle with water, cease respiratory movements and move for-
ward by subtle movements of the suckers. Upon close ap-
proach, in a single motion arms II close the pincer while arms
IV dart forward from underneath. Most commonly the Pincer
Feeding Approach was used when one of the other two at-

44 AMER. MALAC. BULL. 7(1) (1989)

tack maneuvers failed. In any of the attack maneuvers, if the
prey was lost sight of and had escaped, the octopuses would
immediately go to exploratory behavior, extending the arms
in all directions and investigating cracks and the undersides
of rocks.

Octopus briareus would commonly attack, kill and eat
many crabs in succession. With smaller prey, additional crabs
were captured and held in the web until the first was killed
and eaten, at which time the next would be moved to the beak,
killed and consumed. In one demonstration, an adult octopus
captured 50 crabs (Uca sp., 15 mm carapace width). This oc-
topus completely consumed 40 crabs, nine were partially
eaten and one escaped unharmed. A half-grown octopus, 40
mm ML, captured and consumed 17 Uca of the same size.
Thus they have a large appetite.

The exact method of killing the prey is yet unknown.
However, on numerous occasions crabs are seen to tremble
violently from two to four minutes after they are held tightly
near the buccal region of the octopus. The crabs were often
held with their chelipeds towards the buccal mass, and in
these animals no bite marks are found anywhere on the
carapace. It is possible therefore that the toxin from the
posterior salivary gland is released from the buccal area and
absorbed directly through the gills of the crab. Bacq (1951)
described a similar situation and Nixon (1984) has
demonstrated that octopuses can externally digest the
arthrodial membrane and the musculo-skeletal attachments
of crabs without penetrating the exoskeleton. In other cases
the crab is held near the buccal area in the reverse position
and it is possible that the octoups is injecting the toxin through
a small hole in the membranous joint of the carapace (Ghiretti,
1959). This issue requires further study.

REACTIONS TO PREDATORS AND NOXIOUS STIMULI:
Table 5 is asummary of antipredatory adaptations of Octopus
briareus. This table was constructed from laboratory and field
observations, but some of the secondary and tertiary defenses
are speculative. Like most cephalopods O. briareus spends the
majority of its time concealed from predators. They are able
to avoid attracting attention of most foraging predators with
their malleable body form, their nocturnally active cycle and
by remaining motionless against the substrate. Once detected
by a predator, they either flee or use some form of flash
behavior to make a predator hestitate in its attack sequence
(all four acute patterns are used in this type of situation). All
of these reactions have been observed in the laboratory when
an exoerimenier moves a hand swiftly toward an octopus in
jis tank or creates a similar artificial disturbance.

Gruber (1973) observed the reactions of Octopus
briareus to eels (Gymnothorax moringa) in the laboratory. In
249 trials he found the following order or reactions: no
response, 148; Uniform Darkening with papillation, 51; flight
response, 35; inking, 9; Protective posture, 6. His experimental
apparatus was not natural and thus this order of occurrence
can not be construed as the natural response reactions.
However, they do give an idea of the types of reactions an
ociopus is likely to use. In one of our own experiments O.
briareus showed the Deimatic pattern to a moray eel after it

had bitten one of the arms off. In that trial the octopus then
followed the Deimatic pattern with a large discharge of ink
and no further attack occurred.

ACKNOWLEDGMENTS

R. T. Hanlon thanks the members of his M. S. thesis commit-
tee: Dr. Gilbert L. Voss (Chairman), Dr. John R. Southam, Dr. Jon
C. Steiger, Dr. William W. Hay and Dr. Won Tack Yang. V. Ponmatton
and S. Hess helped maintain octopuses reared in Miami in 1973.
Special thanks go to John W. Forsythe who helped with growth
analyses and reared most of the octopuses in the 1981 and 1983
laboratory experiments, which were funded jointly by DHHS Grant
#RR01279 and the Marine Medicine budget of The Marine Biomedical
Institute, The University of Texas Medical Branch at Galveston.

M. R. Wolterding is especially grateful to Francine Spicer, Dr.
Lee Opresko, Barbara Mayo, Richard Schekter and Dr. Norman
Engstrom who helped care for the animals during absences from the
laboratory. He also thanks members of his M. S. committee: Dr. G.
L. Voss (Chairman), Dr. D. Moore, Dr. L. Thomas and Dr. R. Steven-
son. He is especially grateful to Dr. Voss for help throughout graduate
study and for financial support through NSF Grants GV-24030,
GA-11127, GA-1493, GB-5729x, and a grant from the National
Geographic Society.

We are both grateful for a review by Sigurd Boletzky and the
typing assistance of Laura Koppe in Galveston. Finally, we dedicate
this paper to the late Professor Gilbert L. Voss for his support of
students and his many important contributions to cephalopod biology.

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Date of manuscript acceptance: 6 August 1988

THE ECOLOGY OF OCTOPUS BRIAREUS ROBSON IN A
BAHAMIAN SALTWATER LAKE

RICHARD B. ARONSON
DEPARTMENT OF PALEOBIOLOGY
NATIONAL MUSEUM OF NATURAL HISTORY
SMITHSONIAN INSTITUTION
WASHINGTON, D. C. 20560, U. S. A.

ABSTRACT

This paper describes a dense population of Octopus briareus Robson living in Sweetings Pond,
a Saltwater lake on Eleuthera Island, Bahamas. The animals sheltered in cavities within or under discrete
sponge, coral and bivalve formations, and these dens appeared to be limiting. In general, O. briareus
occupied dens for periods on the order of days. They usually remained in their dens during the day,
except for mating and occasional hunting, and foraged primarily at night. They preyed on bivalves,
crabs, fishes, mysid shrimps, polychaetes and each other. Apart from cannibalism, there were no obser-
vations of predation on Sweetings Pond O. briareus.

Copulation was observed three times during the day, and was similar to the laboratory observa-
tions of other investigators. Females guarding eggs were observed year-round in Sweetings Pond, in
contrast to the reproductive seasonality of Octopus briareus off southeastern Florida. Egg and clutch
sizes, development times, the high hatching success (98.8%), and the mean size of brooding females
were similar to previous findings for O. briareus. Most animals showed signs of injuries in the form
of scars, or severed or regenerating arms. Animals suffered injuries in territorial and cannibalistic en-
counters, and during mating.

In contrast to Sweetings Pond, octopuses were rare off the west coast of Eleuthera. Predators,
rather than dens, probably limit Octopus populations in coastal habitats. The high lake density was
due in part to the reduction in number of predatory fishes. Data presented here form a basis for future
comparisons with coastal Octopus populations.

Octopuses exploit a variety of benthic habitats, in-
cluding rocky shores and coral reefs, where they face strong
competition and predation pressure from fishes (Packard,
1972). Octopuses can be important predators themselves in
these habitats (Fotheringham, 1974; Simenstad et al., 1978;
Fawcett, 1984), and they influence the behavior of certain
marine invertebrates (Ross and Boletzky, 1979; Wells, 1980;
Fawcett, 1984). Until recently, however, little was known about
octopus ecology, due primarily to their cryptic lifestyle.

Yarnall (1969) observed Octopus cyanea Gray under
seminatural conditions (in artificial ponds) and describes hunt-
ing behavior, daily activity cycles and a dominance hierarchy
based upon size. The activity cycle of O. vulgaris Cuvier has
been studied (Altman, 1967; Kayes, 1974; Mather, 1988) and
this economically important species has been examined from
a fisheries viewpoint (Hatanaka, 1979a,b; Guerra, 1981; Smale
and Buchan, 1981). The ecology of O. dofleini (Wulker) is the
subject of ongoing research in British Columbia (Hartwick et
al., 1978a, b, 1981, 1988; Hartwick and Thorarinsson,

1978), and ecological field studies of O. joubini Robson (But-
terworth, 1982; Mather, 1982) and O. bimaculatus Verrill (Am-
brose, 1982, 1988) have been carried out as well.

From 1980 to 1983, | examined the ecology of Octopus
briareus Robson in Sweetings Pond, a saltwater lake in the
Bahamas. The absence of predatory fishes in Sweetings Pond
has resulted in a unique community, of which O. briareus is
the top carnivore (Aronson and Harms, 1985; Aronson, 1986).
Here | treat a variety of topics pertaining to O. briareus ecology
and behavior and, where information is available, compare
the results to previous octopus studies. Hopefully, this paper
will serve as the basis for future comparison with octopus
populations in coastal habitats.

STUDY AREAS

Sweetings Pond is situated at the north end of
Eleuthera Island, Bahamas (25921’N, 76°30’W; Fig. 1). It is
surrounded by karstic limestone and has surface area of

American Malacological Bulletin, Vol. 7(1) (1989):47-56

47

48 AMER. MALAC. BULL. 7(1) (1989)

0.92 km2. The maximum depth is 15.3 m. Unconsolidated sedi-
ment covers a limestone pavement forming the bottom of the
lake. The water chemistry and tidal cycle of Sweetings Pond
indicate a connection to the west coast of Eleuthera via one
or more restricted subterranean passages and/or percolation
through the porous rock (Aronson, 1985). Human activity is
negligible.

Figure 2 illustrates the marked benthic zonation ob-
served in the area of the Cove Entry (Fig. 1) in 1980. From
shore to a depth of approximately 2 m, thick, fluffy mats of
the filamentous green alga Cladophora crystallina (Roth)
dominated the substratum. Between 2 and 8 m depth, sponge,
coral and bivalve formations (Table 1) were scattered over the
bottom. These structures either rested atop the sediment or
were loosely buried. Because of the discrete nature of the

20 km

Atlantic Ocean

vernor s Harbour

Eleuthera Bank

Rock Sound

Fig. 1. Map of Eleuthera Island, Bahamas, showing location of
Sweetings Pond. Inset: map of Sweetings Pond, showing locations
of Study Plots (numbered squares). R11 is the survey, collecting and
experimental area. Dashed lines are dirt tracks. Adapted with per-
mission from Bahamas Department of Lands and Surveys maps.

Algal Mat

formations, this zone was termed the ‘‘patch zone.” The main
concentration of Octopus briareus occurred in cavities in and
under some types of patch zone formations. Other octopuses
were found under limestone rocks at shore. The patch zone
thinned at its deep end, being composed mostly of flat orange
sponges at a depth of 7.5 m. From the end of the patch zone
to the center of the lake, the bottom consisted mainly of bare
sediment, with scattered clumps of algae.

Notable in the patch zone was the high density of
ophiuroids, particularly the epifaunal suspension-feeder
Ophiothrix oerstedi Lutken, which occurred at densities up to
434 ind./m?2 (Aronson and Harms, 1985). Other conspicuous
mobile invertebrates were the large spider crab Mithrax
spinosissimus (Lamarck), the starfish Echinaster sentus (Say),
the sea urchin Echinometra viridis Agassiz, the polychaete
Eunice rubra Grube and the gastropod Fasciolaria tulipa
(Linnaeus).

In 1982 and 1983, much of Sweetings Pond did not
follow the generalized profile of figure 2. Considerable areas
were overgrown by Cladophora mats, and these mats expand-
ed and regressed during 17 months of study. The cove that
contains the Dock Entry (Fig. 1) was entirely patch zone in
1980. In February, 1983, almost the whole cove was covered
by Cladophora, but the mats were dying back by June, 1983.
Off the Cove Entry, a major portion of patch zone was covered
by algal mat in 1982 to 1983. The algae destroyed all patch
zone formations, leaving bleached coral skeletons and empty
shells of Chione cancellata (Linnaeus) (the most common in-
faunal bivalve) and Arca imbricata Brugiére after it regressed.
In this way much Octopus habitat was destroyed, including
Study Plot 1 (Fig. 1). The cause of these dramatic changes
in algal cover is unclear, but nutrient input via runoff from the
cultivated fields surrounding Sweetings Pond could be
responsible.

The fish fauna of Sweetings Pond was remarkably
depauperate: 17 species from 15 families were recorded (Aron-
son and Harms, 1985). By contrast, 126 species from 50
families were recorded in shallow water (<6 m depth) off the
west coast of Eleuthera in the vicinity of Sweetings Pond. The
only potential predators of Octopus sighted in hundreds of
hours of diving in the lake were five schoolmasters, Lutjanus
apodus (Walbaum), one Nassau grouper, Epinephelus striatus
(Bloch), and one moray eel, Gymnothorax funebris Ranzani
(see Randall, 1967).

Depth (m)

Bare Sediment

Scattered Algal Mat

SS ee
50m

Fig. 2. Benthic profile from the Cove Entry to the center of Sweetings Pond in July, 1980.

ARONSON: OCTOPUS ECOLOGY 49

Table 1. Principal formations in the patch zone of Sweetings Pond. Combinations of these types were

also encountered.

TAXA ASSIGNED NAME

A. Sponges
Xestospongia sp.
Halicometes sp.
Suberites sp.
Reniera sp.

B. Coral-dominated
Porites porites (Pallas)
Arca imbricata Brugiére
Porites astreoides Lamarck
Siderastrea spp.

C. Bivalve-dominated
Arca imbricata
Chama macerophylla (Gmelin)
Lima scabra var. tenera (Sowerby)
Pinctada imcricata Roding

brown sponge
orange sponge #1
orange sponge #2
white sponge

bivalve clump

REMARKS

often with Arca attached

P. porites-bivalves

large head with scattered Arca
small heads; rare

chiefly Arca and Chama

METHODS

Ecological information was collected in 1980 and 1982-83
during approximately 450 SCUBA and snorkel dives from the
shore of Sweetings Pond. For comparison, more than 150 day
and night dives were made off the west coast of Eleuthera
between Governor’s Harbour and the Glass Window (Fig. 1).

The discrete nature of the patch zone formations
facilitated mapping, and their lack of strong attachment to the
substrata allowed them to be turned over carefully and then
replaced. In July 1980, three surveys were conducted in a
30x30 m area of the patch zone (Study Plot 0; Fig. 1), gridded
with nylon line to form squares 3 m on a side. The surveys
involved examining each formation in the plot for Octopus
briareus. When an individual was found, it was captured,
measured, and sexed; and the formation position, type, size,
and depth were recorded. The animal was then returned to
its den. In addition, numerous census dives were made in
the patch zone to the southeast of the Cove Entry. Two divers
swam zig-zag patterns through the entire depth range of the
patch zone and captured each animal encountered. The same
ecological information was taken as in the surveys of Study
Plot 0, but the position of the formation was not mapped.

Two 27x30 m grids (Study Plots 1 and 2), with the same
element size as Study Plot 0, were constructed in the patch
zone in April 1982. These were surveyed monthly. Study Plot
1 was not surveyed after July 1982 because it became almost
completely overgrown with Cladophora, whereas Study Plot
2 was surveyed through July 1983. Monthly patch zone survey
dives were also made off the Cove Entry and southeast, near
Study Plot 2. These dives were made as close in time as
possible (usually on the same day) to the Study Plot surveys,
to minimize the possibility of encountering individual Octopus
briareus twice. The divers’ search procedures were the same
as for the census dives of July 1980.

All research on den ecology was carried out between
the hours of 0800 and 1300 EST. During this time interval,
96% (457/476) of the Octopus briareus encountered in

Sweetings Pond under natural conditions were in dens (see
Diel Activity Pattern). Dives were made at all hours of the day
and night to observe diel activity patterns, feeding habits and
reproductive behavior.

For convenience, the word ‘‘dorsal’’ will be used to refer
to the upper surface of the octopus when it is resting on the
bottom or swimming in its usual orientation. The designation
“ventral” will be in keeping with the above definition. This
simplification follows Wells’ (1978) suggestion.

RESULTS AND DISCUSSION

DENSITY, SPATIAL DISTRIBUTION
AND DEN ECOLOGY

Octopus briareus occurred at high density in Sweetings
Pond. In July 1980 the three surveys of Study Plot 0 (900 m2)
yielded counts of 11, 14 and 16 individuals (mean 15.3 + 3.1
SD ind./1000 m2). Density varied from area to area of the patch
zone, apparently depending on the availability of suitable dens
(see below). In April, 1982, for example, Study Plots 1 and
2 (810 m2) were surveyed and found to contain 12.3 + 1.2
SD and 4.1 + 1.5 SD ind./1000 m2, respectively (n = 3
surveys). The 1982-83 mean for Study Plot 2 was 7.9 + 36
SD ind./1000 m2 (Aronson, 1986).

A nearest neighbor analysis (Clark and Evans, 1954;
Poole, 1974) was performed on the spatial distribution of
Octopus briareus in the three surveys of Study Plot 0. There
was no significant deviation of mean nearest neighbor
distance from that expected had the animals been dispersed
randomly. The same analysis was performed for four
categories of dens potentially occupied by O. briareus: brown
sponges showed significant clumping (p<0.005), Porites
porites-bivalve clumps were evenly dispersed (p< 0.0005),
and bivalve clumps and orange sponges were distributed ran-
domly (p>0.40).

While it was impossible to predict whether a particular
formation was suitable as a den, some formations were oc-
cupied frequently while others remained unoccupied (Fig. 3;

50 AMER. MALAC. BULL. 7(1) (1989)

see also Aronson, 1986 for statistical analysis). The implica- were limiting there (Tauchi and Matsumoto, fide Mottet, 1975).
tion is that dens could be the limiting resource for O. briareus Ambrose (1982) suggests that dens are more likely to be
in Sweetings Pond. This hypothesis is supported by ex- limiting in soft-bottom communities like Sweetings Pond. In
periments in which local density was increased substantially general, rocky subtidal habitats contain more crevices, but
by adding artificial dens to patch zone plots (Aronson, 1986). how cavity-occupying fishes and invertebrates affect the
Mather (1982) found that the distribution of Octopus availability of dens is unknown (Aronson, 1986).
joubini in a Florida soft-bottom community correlated with the Hartwick et a/. (1978a) found that the Octopus dofleini
availability of dens. Ambrose (1982), on the other hand, con- in their study were more evenly spaced than expected in a
cluded that suitable dens were not limiting to O. bimaculatus random distribution. This result, combined with an observa-
on hard substrata off Santa Catalina Island, California. In a tion of intraspecific fighting over a den (Kyte and Courtenay,
removal experiment, Hartwick et a/. (1978a) found that some 1977), helps make the case for the commonly held view that
dens were occupied by O. dofleini of similar size to the O. dofleini is territorial (Hartwick et a/., 1978a). Woods (1965)
previously evicted occupants; however, dens were not limiting and Cousteau and Diolé (1973) present anecdotal evidence
in the rocky subtidal off Vancouver Island, British Columbia for territoriality in natural populations of O. vu/garis; however,
(Hartwick et a/., 1988). On the other hand, cavities may have Altman (1967) and Kayes (1974) found no evidence of territorial
been limiting in offshore, soft-bottom habitats (Hartwick et al., behavior in this species. At Kayes’ (1974) field site off Malta,
1988). Adding artificial dens to an octopus fishing ground off dens ‘‘were found wherever the substrate was suitable, and
Japan increased the catch dramatically, implying that dens occasionally occupied holes were only 1 m apart.’ Guerra
| 2 3 6 ¢ 8 ,
fe)
$ ° Sse .
J ea fe)
DB : 0
| O
2 os |
ge O | é le ve th
H rele < p
o |
8 +
: |
G A 0 Oo (o)
°6
: A
®
F la
et le 4 Im
A \@
a [ ela = A
A A
| | =
D : al | 46 aa O
e Zn
a c ato | 4
° A
C e io! e a!
e
®
| =a z
2 °
B | e | :
A
-e : J <
|
A | Ola A |
4 | 2 ee! A i
A «99 |
j
Fig. 3. Map of Study Plot 2, showing locations of Octopus briareus dens mapped during 1982-83. The number next to a formation is the number — |
of different O. briareus that were found in or under the formation. Solid triangle, Porites astreoides; solid circle, brown sponge; open circle,
orange sponge; solid square, mixed clump of FP. porites and bivalves; open square, bivalve clump; open triangle, other. Formation types that |
were never occupied are not mapped, and the only ‘‘other’’ formations mapped are those that contained O. briareus.

ARONSON: OCTOPUS ECOLOGY 51

(1981) concluded that O. vulgaris off west Africa were dis-
persed randomly within patches, the patches being of vary-
ing density. The minimum nearest neighbor distance observed
under natural conditions in the present study was 80 cm, for
a female and a juvenile inhabiting brown sponges. In artificial
den experiments, O. briareus pairs of the same and opposite
sexes occupied polyvinyl chloride (PVC) tubes as close as
15 cm (Aronson, 1986). Considering the spacing of natural
dens seen in the Study Plots (Fig. 3), den defense probably
did not affect nearest neighbor distance. Suitable dens were
simply too far apart.

TENURE OF DEN OCCUPATION

The maximum tenure of den occupation in Sweetings
Pond (based on daily den checks in the morning) was = 25
days for animals not brooding eggs. This number is derived
from observations of a female (mantle length 8.0 cm) that oc-
cupied the cavity under a brown sponge, disappearing after
25 days of observation. She had a swollen gonad and was
apparently about to lay eggs, which could account for the
length of her stay. Incidental observations of Octopus briareus
denning under limestone rubble along the shore just off the
Cove Entry gave a tenure of occupation on the order of a few
days. Artificial dens (PVC tubes; Aronson, 1986) were also oc-
cupied for one to a few days, with a maximum of seven days
(n = 81 observations).

The 1982-83 monthly surveys support these results.
Eighty-five of 93 Octopus briareus were not found in the same
den the previous month. Of the eight remaining, three were
egg-brooding females seen in two consecutive months, one
was an adult male seen in two consecutive months (recog-
nized by injuries) and four were ambiguous cases (they might
have been the same animals as in the previous monthly
survey). The male was found twice in the same den 35 days
apart.

Altman (1967) states that Octopus vulgaris were found
in “‘permanent” or ‘‘temporary’’ homes in the field. ‘‘Perma-
nent’’ homes were occupied for at least two consecutive days;
of 37 dens observed, four were occupied for the duration of
the study (25 days). Unfortunately, no information is given on
the sexual state of these animals. Most O. vulgaris occupied
dens for one to two days in Kayes’ (1974) field study. Yarnall
(1969) reported a maximum tenure of den occupation of 23
days for O. cyanea in artificial ponds and Van Heukelem (1966)
reported a maximum occupancy of 35 days in the field
(Hawaii). Most O. dofleini in the study by Hartwick et al. (1984)
were found in the same dens for at least one month. Their
results ‘‘indicate a pattern of large-scale movement inter-
spersed with periods of residence in a relatively small area...’
By far the longest periods of occupation observed are for O.
bimaculatus. Almost half the population examined by Ambrose
(1982) occupied the same dens for more than one month, and
three individuals remained in the same dens for more than
five months. O. briareus appears to be a fairly mobile species,
both in Sweetings Pond and in coral reef habitats (Hochberg
and Couch, 1971; J. Wodinsky, pers. comm.).

DEN BLOCKING AND EXCAVATION
Octopus briareus frequently blocked the entrances to

their dens with small pieces of bivalve clump (mostly Arca
imbricata), empty bivalve shells, pieces of live or dead Porites
porites and live Chione cancellata. Such acitivity was most
obvious when the animals resided in PVC or acrylic tubes but
was sometimes unambiguously the case with natural dens
as well. Blocking is a well-known behavior in O. vulgaris
(Legac, 1969; Cousteau and Diolé, 1973) and also occurs in
other species (Van Heukelem, 1966). The separate topic of
middens outside octopus dens will be considered in Diet.

Den excavation has been reported in the literature for
a number of species (Yarnall, 1969 for Octopus cyanea;
Hochberg and Couch, 1971 for O. macropus Risso; Cousteau
and Diolé, 1973 for O. vulgaris; Hartwick et al., 1978a for O.
dofleini; Ambrose, 1982 for O. bimaculatus). Excavation by fun-
nel blasts from O. briareus was observed once in Sweetings
Pond, and excavations under patch zone formations were in
many cases obvious because of the different color of the sand
that had been exposed.

DIEL ACTIVITY PATTERN

Hochberg and Couch (1971) characterize Octopus
briareus as a nocturnal hunter, based on field observations
in the United States Virgin Islands. Hanlon’s (1975) field obser-
vations from a number of Caribbean localities support their
conclusion. Ninety-six percent of O. briareus (n = 476) in
Sweetings Pond were found in dens during the morning
(0800-1300 hours EST). By contrast, 94% (n = 35) of in-
dividuals at night were out in the patch zone (x2=290.80, df=1,
p<0.005). Of these, four were sitting atop formations (their
dens?) and 29 were moving along the substratum. Three out
of 20 animals captured at night had prey (see Diet), although
one individual captured during the day was carrying two small
spider crabs, Pitho sp. (undescribed). The three observed in-
stances of copulation occurred during the day (see Reproduc-
tion: Mating Behavior).

Different activity cycles have been reported for different
Octopus species. Octopus vulgaris makes long excursions at
night and early in the morning, with short trips during the day
(Altman, 1967; Kayes, 1974; see also Mather, 1988). Altman
(1967) raises the possibility that the activity pattern of O.
vulgaris is related to those of prey or predator species. The
prey activity hypothesis could also apply to O. briareus in
Sweetings Pond: the bivalve Laevicardium laevigatum (Lin-
naeus) comes out of the sediment at night (see Diet). O. joubini
is nocturnal (Mather, 1982, 1984), whereas O. dofleini showed
only a slight activity peak at night in a sonic tagging study
(Mather et a/., 1985). Houck (1982) demonstrated that three
sympatric Octopus species in Hawaii displayed differences
in diel activity cycles, possibly reducing competition or preda-
tion on smaller Octopus species by larger ones. The isolated
population of O. briareus in Sweetings Pond would be ideal
for comparison with coastal populations in a study of
behavioral character displacement.

DIET

Information on the feeding habits of Octopus spp. has
been derived primarily from the examination of piles of prey
remains, or middens, that the animals leave outside their dens

52 AMER. MALAC. BULL. 7(1) (1989)

(Van Heukelem, 1966; Altman, 1967; Hochberg and Couch,
1971; Kayes, 1974; Hartwick et a/., 1981; Smale and Buchan,
1981; Ambrose and Nelson, 1983). Obviously, such middens
only preserve information on prey items that have hard parts
and that are consumed in the den (Smale and Buchan, 1981).
Furthermore, prey discards can disappear from middens due
to biotic (hermit crabs) or abiotic (currents and surge)
taphonomic processes (Ambrose, 1983). Middens have re-
vealed that crustaceans and mollusks are important consti-
tuents of the diet of octopuses, but the information loss
associated with middens argues against their use in quan-
titative analyses.

Octopus briareus middens were uncommon in the
patch zone of Sweetings Pond, possibly, as Wolterding (1971)
suggests, because of the high mobility of this species. At shore
just off the Cove Entry, a number of middens were found out-
side occupied dens. Collections revealed the following prey
species: the bivalves Laevicardium laevigatum, Brachydontes
domingensis (Lamarck) and Chione cancellata, and the crab,
Pitho. This species list is based on fresh discards (i.e. no signs
of pitting or algal growth on them). The bivalves Lima scabra
var. tenera, Pinctada imbricata, Codakia orbiculata (Montagu),
Polymesoda maritima (d’Orbigny) and Chama macerophylla
were also found in middens and in dens, but it was difficult
to determine the freshness of these shells. C. cancellata, the
most common infaunal bivalve in Sweetings Pond, presents
a problem. Empty valves of this species are used for block-
ing by O. briareus, as are living individuals. O. briareus did
not eat bivalves in a laboratory study (Wolterding, 1971).

An Octopus briareus captured during the day was carry-

ing two Pitho sp. and an octopus encountered during a night
dive held a mysid shrimp. O. briareus also eat fishes and
polychaetes (Hochberg and Couch, 1971; Wolterding, 1971;
Hanlon, 1975). One octopus, captured at night, was eating
the fish Callionymus pauciradiatus Gill, and another examined
at night was holding an unidentified polychaete. On two oc-
casions, O. briareus found in their patch zone dens during
the day were eating polychaetes, Eunice rubra.

Cannibalism is known in field populations of Octopus
vulgaris (Smale and Buchan, 1981), O. dofleini (Hartwick et
al., 1978a) and O. bimaculatus (Ambrose, 1984), and this
behavior is well documented for a variety of species kept in
aquaria (e.g. Lane, 1974). Wolterding (1971) and Hanlon and
Forsythe (1985) observed cannibalism by O. briareus in the

Table 2. Instances of cannibalism by Octopus briareus in Sweetings
Pond. All observations are from 1982 (?, sex indeterminable in the
field; —, no data).

CANNIBAL PREY
Mantle Mantle
Date Length (cm) Sex Length (cm) Sex
13 Apr 43 M 18 F
17 Apr 3.0 ts 2.0 M
11 May 7.0 F = =
13 July 8.5 F 53 M
17 July 7.0 F 55 —
19 Aug 5.5 M 28 F

laboratory, and Hanlon (1983) discusses some of the instances
observed in the present study. Six observations of cannibalism
were made in Sweetings Pond in 1982-83 (Table 2). In the
observation of 13 July 1982, the female copulated a few
minutes after she was discovered eating a different male.
Direct observations and examination of crop contents revealed
that the arms are eaten first. One victim (17 July 1982) had
had all of its arms eaten yet was still alive. When removed
from the grasp of the cannibalizing female, it made weak at-
tempts to swim and crawl away. The potential interacton be-
tween cannibalistic and mating motivational states is dis-
cussed in the next section.

REPRODUCTION: MATING BEHAVIOR

Three copulations were observed. One occurred in July
1982 and is discussed by Hanlon (1983). This mating involved
a 5.3. cm mantle length male and an 8.5 cm female, and lasted
approximately 60 min. The second copulation, in August 1982,
involved a 6.0 cm male and a 7.0 cm female. The female had
been in an artificial den placed in the patch zone at 3.7 m
depth. This den consisted of a length of 10 cm diameter acrylic
tubing with a 5 cm entrance. The male was hiding under sand-
bags that covered the den. | removed the sandbags and
evicted the female, at which point the male instantly leapt upon
her dorsal mantle and they copulated. Mating lasted 67
minutes. At the end of copulation, the male dismounted
abruptly and swam away (the male in the first copulation did
not leave as quickly). The female was left with about 40 cir-
cular sucker scars on the posterior end of her dorsal mantle.
The third instance was observed briefly in January 1983, but
extensive observations were not possible (C. A. Harms, pers.
comm.).

In all cases, mating was similar to laboratory observa-
tions (Wolterding, 1971; Hanlon, 1975) and occurred during
the day. Noteworthy is the abruptness with which the male
left the female in the second case. Females could attempt
to cannibalize males after mating. In the first case, mating
occurred just after the female had cannibalized a male
(Hanlon, 1983). By retreating hastily, the male in the second
copulation could have been avoiding capture by the female;
in fact, he was already missing all of his third left and part
of his first right arms. As mentioned in Injuries, the only causes
of arm loss in Sweetings Pond appear to be intraspecific
fighting and cannibalism.

The absence of precopulatory behavior in Octopus
briareus under laboratory and field conditions (Hanlon, 1983
and references therein; this study) is in marked contrast to
the ritualization seen in O. cyanea (Van Heukelem, 1970; Wells
and Wells, 1972). Male O. vulgaris perform a “‘sucker display”’
when initiating mating with a larger female (Packard, 1961).
It could be that the pattern of physical contact itself (i.e. the
male’s climbing on top of the female’s mantle) serves to iden-
tify the Octopus male to the female (Wells and Wells, 1972).
Considering the variation in copulatory behavior reported in
O. cyanea and O. vulgaris (Wells and Wells, 1972; Wodinsky,
1973; also see Mather 1978 on O. joubini), including instances
in which the males of these species also leapt upon the
females, the apparent simplicity of mating initiation in O.

ARONSON: OCTOPUS ECOLOGY 53

briareus merits further attention.

REPRODUCTION: EGGS AND EGG BROODING

In censuses from March 1982 to July 1983, females
guarding eggs were recorded in Sweetings Pond in each
month except June and July 1983, with annual peaks in
February and March (Aronson, 1986). In southeastern Florida,
Octopus briareus seem to display a more strongly seasonal
maturation and breeding cycle (Hanlon, 1983); the degree of
synchrony may therefore vary from population to population.

The mean egg length, computed from eggs at develop-
ment Stage IV (Wells and Wells, 1977) or earlier was 10.50
+ 068 SD mm (range 9.0 - 12.0 mm, n = 99; based on 11
eggs each from nine females). The mean mantle length of
females guarding eggs was 6.1 + 0.92 SDcm(n = 25). Two
egg broods, followed from stage IV or earlier to hatching, gave
a development time range of 50-67 days. The first female
brooded her eggs for 50 days (11 May-30 June 1982;
23.5-32.0°C) and the second for 67 days (18 January-26 March
1983; 20.5-23.5°C). The value of 36 days quoted by Hanlon
(1983) for development time in Sweetings Pond was an early
minimum estimate.

Clutch sizes were determined from egg strands col-
lected within two days of hatching. For each egg strand, the
number of empty attached capsules and the number of egg
stalks from which the capsule had been separated were
counted. This procedure avoided the destruction of clutches,
which would have been necessary had the eggs been counted
prior to hatching. The error caused by possible loss of hatched
strands was minimal. The mean clutch size was 267.3 + 99.2
SD eggs (range 97 - 414 eggs; n = 8). There was no correla-
tion between clutch size and the mantle length of the brooding
female (product-moment correlation coefficient, r = —0.20,
n = 8, p > 0.90; Sokal and Rohlf, 1969). Hatching success
was high: 1453 of 1471 eggs (from 6 females) hatched, for
a success rate of 98.8%. The egg sizes, clutch sizes, develop-
ment times, hatching success and average size of brooding
females were similar to previous findings for Octopus briareus
(Hanlon, 1983).

INJURIES

Two classes of injury were noted for Octopus briareus
in Sweetings Pond: arm injuries and scars. Arm injuries con-
sisted of severed or regenerating arms. Arm injuries can result
from cannibalism, and O. briareus can also lose or autotomize
(Lange, 1920) arms in fights. In a fight staged by divers be-
tween a female of mantle length 4.0 cm and an 8.0 cm female,
the smaller one lost three arms. However, neither octopus lost
any arms in the only fight observed under natural conditions.
In this case two males, 4.5 and 5.5 cm in mantle length, were
struggling underneath a brown sponge. The larger individual
clearly had the advantage in both fights. It was impossible
to tell whether these were territorial or predator-prey en-
counters, or both.

Scars occurred in a number of forms (Fig. 4). They
were usually on the head, or dorsal or ventral mantle, although
they occurred on the arms as well. Many scars were rings.
In other cases, white lesions of varying sizes appeared on

Fig. 4. Sample of Octopus briareus dorsal mantle scar patterns. All
scars are drawn in negative. A, sucker marks and a triangular skin
lesion; B, sucker marks and lines; C, sucker marks, perhaps from
a single O. briareus arm; D, extensive skin lesions.

the mantle, arms or webbing where patches of skin had been
removed. Still other scars were dark or light lines.

From the second observation of copulation, it is ob-
vious that the dark ring scars were sucker marks. Skin lesions
could arise from the suckers or beak bites of other octopuses;
they could also result from infections or from scraping against
rough surfaces. Linear scars, rarer than the other two
categories, are of unknown origin. Scars can be acquired dur-
ing mating, in encounters with cannibals and during fights.

There was no evidence of injuries caused by in-
terspecific interactions (in contrast to Hartwick et a/., 1988).
The only animals common enough and large enough to have
posed a threat to Octopus briareus were spider crabs, Mithrax
spinosissimus, which reached at least 20 cm in carapace
width. However, these herbivorous crabs became quite
alarmed and retreated when O. briareus were placed near
them.

Larger Octopus briareus displayed the results of injuries
more frequently than did smaller ones (Table 3). The most
obvious explanation is that the probability of an older in-
dividual having interacted with a conspecific is greater. Males
and females not guarding eggs showed about the same fre-
quency of injury. Females guarding eggs showed a high rate
of arm loss and scarring, possibly due to cannibalism by
copulating males and the effects of their suckers, and to the
general deterioration associated with egg-brooding. It would
be interesting to see whether O. briareus in low-density coastal
populations, which presumably encounter conspecifics less

54 AMER. MALAC. BULL. 7(1) (1989)

Table 3. Breakdown of injuries for Octopus briareus in all 1982-83
surveys combined. ‘‘Unsexable’’ individuals were those <4.0 cm
mantle length.

A. Scars
Number of animals (proportion)

Sex category With injury Without injury N

Males 79 (0.73) 29 (0.27) 108
Females (no eggs) 62 (0.77) 19 (0.23) 81
Females guarding eggs 14 (1.0) 0 (0) 14
Unsexed Adults 4 (1.0) 0 (0) 4
Unsexable 10 (0.12) 73 (0.88) 83

B. Severed or regenerating arms, or parts of arms
Number of animals (proportion)

Missing Missing

arm or only Without
Sex category part tip(s)@ injury N
Males 11 (0.10) 9(008) 89(082) 109
Females (no eggs) 11 (0.13) 0 (0) 73 (0.87) 84
Females guarding eggs 6 (055) 0 (0) 5 (0.45) 11
Unsexed Adults 5 (1.0) 0 (0) 0 (0) 5
Unsexable 2 (0.02) 1 (0.01) 85 (0.97) 88

aThe distal centimeter or less of the arm.

often, show lower injury frequencies.

It was possible to identify individual animals by a com-
bination of size (good on a short-term basis), sex, mantle scar
pattern, and arm injury/regeneration pattern. Individual iden-
tifications were used in mapping the occupants of Study Plot
2 and in determining the tenure of den occupation. The scar
patterns were very distinctive, and although widely divergent
scar patterns were chosen for figure 4, far more subtle distinc-
tions could be made. Cephalopod arms take on the order of
months to regenerate (Lange, 1920; Féral, 1978). While this
is not as sure an identification technique as branding (Van
Heukelem, 1973) or subcutaneous dye injection (Altman, 1967;
Hochberg and Couch, 1971; Kayes, 1974), it eliminates the
trauma of those procedures. To compensate for the possible
loss in accuracy of animal identification, ambiguous cases
were always rejected as data. The technique obviously worked
better for larger than for smaller animals.

OCTOPODS OFF THE COAST OF
ELEUTHERA ISLAND

Octopus briareus occurred in and around subtidal
crevices along the rocky west coast of Eleuthera. Compared
to the density in Sweetings Pond, Octopus spp. were rare off
the west coast. In 1982 and 1983, Octopus were seen four
times in shallow diving (<5 m) at sites from Governor’s Har-
bour to the Glass Window. One O. macropus was seen at
night, two O. briareus were observed, one at night and one
in the afternoon, and an Octopus sp. (probably briareus) was
seen in the evening. While it is true that many cavities in the
limestone rock were too deep to be searched effectively, 166
day and night dives were made specifically to search for oc-
topuses. Dens did not appear to be limiting off the west coast
(Aronson, 1986).

GENERAL DISCUSSION AND CONCLUSIONS

The population of Octopus briareus in Sweetings Pond
has persisted from at least 1972 to 1988 (pers. obs.). How the
species was introduced to this ‘‘island’’ is undetermined.
Perhaps introduction occurred from the west coast of
Eleuthera through subterranean passages.

Island populations are often subject to less predation
pressure than they experience on the ‘‘mainland’’, resulting
in high densities (MacArthur and Wilson, 1967). Release from
predation probably accounts for the high density of Octopus
briareus in Sweetings Pond. Of the 29 species of Caribbean
reef fishes listed by Randall (1967) as eating cephalopods,
only two (Epinephelus striatus and Lutjanus apodus) were
observed in Sweetings Pond, and these were rare. A single
moray eel, Gymnothorax funebris, was sighted in the lake. By
contrast, 19 of Randall’s (1967) cephalopod predators occur
off the west coast of Eleuthera and | suspect that predatory
fishes limit Octopus populations there. The low abundance
and diversity of predatory fishes in Sweetings Pond could be
due to chance factors associated with colonization, or the lake
could be too small to sustain populations of top-level carni-
vores (MacArthur and Wilson, 1967). Hamner (1982) has anec-
dotally reported similar island effects for the faunas of the
Palau salt lakes.

In a soft-bottom habitat such as Sweetings Pond, it is
not surprising that dens were limiting to a high-density Octo-
pus briareus population. Predation is probably more impor-
tant than den availability in limiting Bahamian coastal Octopus
populations. The ethological consequences of this ecological
difference are as yet unknown. Whether predators of O.
briareus exert a limiting influence at the recruitment and
juvenile stages (Ambrose, 1988) or by preying on adults is also
not known.

| have argued elsewhere (Aronson and Harms, 1985;
Aronson and Sues, 1987) that Sweetings Pond, which is
dominated by an epifaunal, suspension-feeding echinoderm
(the ophiuroid Ophiothrix oerstedi) and a_ predatory
cephalopod, can be viewed as a modern-day analogue of cer-
tain Paleozoic and Early Mesozoic soft-bottom communities.
The exclusion of predatory reef fishes is apparently respon-
sible for the high density of both species in the lake. Ancient
epifaunal, suspension-feeding communities, populated by
carnivorous ectocochliate cephalopods, could also have per-
sisted due to low predation pressure from fishes (Aronson and
Sues, 1987). Comparing a dense, isolated Octopus briareus
population, and the fauna of Sweetings Pond as a whole, to
coastal populations could provide clues to the structure of an-
cient benthic marine communities.

ACKNOWLEDGMENTS

| thank C. A. Harms, M. E. Day and E. M. Pollak, Jr. for their
assistance in the field. W. H. Bossert, T. J. Givnish, R. T. Hanlon,
L. S. Kaufman, K. P. Sebens and R. D. Turner gave invaluable ad-
vice throughout this study. | also thank A. Brussel, R. R. Olson, M.
R. Patterson, J. Wodinsky and R. M. Woollacott for their comments
and suggestions. The following people kindly identified organisms:
R. T. Abbott, C. J. Bird, K. Fauchald, G. L. Hendler, L. S. Kaufman, R.

ARONSON: OCTOPUS ECOLOGY 55

B. Manning, K. Ruetzler and R. L. Turner. G. C. Mattran of the U.
S. Embassy in Nassau and D. Thompson of Gregory Town, Eleuthera
provided logistical support. R. J. Mooi drafted figures 1, 2 and 4.

This research was funded by a Fulbright Grant, a National
Science Foundation Dissertation Improvement Award (grant
DEB-8114684), an NSF Graduate Fellowship, and grants from the
Lerner-Gray Fund of the American Museum of Natural History, the
Richmond Fund of Harvard University, Sigma Xi, the Explorers Club
and the Hawaiian Malacological Society. Field work was carried out
under a permit from the Department of Fisheries, Commonwealth
of the Bahamas. This paper is based on a doctoral dissertation sub-
mitted to the Department of Organismic and Evolutionary Biology,
Harvard University.

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Ambrose, R. F. 1982. Shelter utilization by the molluscan cephalopod
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Ambrose, R. F. 1983. Midden formation by octopuses: the role of biotic
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Ambrose, R. F. 1984. Food preferences, prey availability, and the diet
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Ambrose, R. F. 1988. Population dynamics of Octopus bimaculatus:
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Ambrose, R. F. and B. V. Nelson. 1983. Predation by Octopus vulgaris
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Date of manuscript acceptance: 28 November 1988

AN ATLANTIC MOLLUSCAN ASSEMBLAGE DOMINATED BY TWO
SPECIES OF CRASSINELLA (BIVALVIA: CRASSATELLIDAE)

WILLIAM G. LYONS
FLORIDA DEPARTMENT OF NATURAL RESOURCES
ST. PETERSBURG, FLORIDA 33701, U. S.A.

ABSTRACT

A total of 136 molluscan species were obtained in benthic grab samples during 12 bimonthly
periods (Sept 1971 - July 1973) at five stations (depths 7-11 m) near Hutchinson Island, east central
Florida; 33 characteristic species constituted 90% of the 4135 specimens. Species distributions were
influenced strongly by sediment composition. Compacted fine and very fine sands of the beach ter-
race supported few mollusks. Well-sorted medium sands supported a small but abundant species group
at an offshore shoal, and two larger species groups were associated with coarse sands and with large
shell particles that entrapped mud and silt in a trough between the shoal and terrace. Two bivalve
species were numerically dominant. Crassinella lunulata (Conrad, 1834) contributed 33% of all specimens
and occurred among large shell particles in the trough; C. dupliniana (Dall, 1903) contributed 14%
of all specimens and favored medium sands at the shoal. C. dupliniana, originally described as a fossil,
has not been recorded previously among living fauna.

Only two studies have examined the species composi-
tion and relative abundance of small mollusks associated with
microhabitats of the continental shelf of the southeastern
United States. Both studies addressed assemblages
dominated by corals (McCloskey, 1970; Reed and Mikkelsen,
1987), so, expectedly, most of the mollusks were species with
affinity for hard substrata. Consequently, very little is known
about species associated with various sediment regimes of
the shelf.

Opportunity to acquire information on sand-bottom
species associations on the inner continental shelf of eastern
central Florida occurred during 1971 through 1973, when the
Florida Department of Natural Resources (FDNR) conducted
a study to assess potential environmental impacts from a
nuclear power plant then under construction at Hutchinson
Island. Offshore environments were sampled with trawls,
plankton nets, and benthic grabs; the surf zone was sampled
with beach seines; and specimens were collected by hand
from a rocky shore community. Several technical reports on
environmental parameters, species composition, distribution,
and seasonal fluctuations of biota from those samples have
been published in Florida Marine Research Publications.

Two species of the bivalve genus Crassinella were
numerically dominant among mollusks in quantitative grab
samples obtained off Hutchinson Island. Associations of those
species with other small infaunal-epifaunal mollusks and the
relationship of those associations to different sediment
regimes are reported here. One of the species, not recognized

previously in Recent fauna, is illustrated and compared with
congeners.

METHODS

Five benthic stations on the nearshore continental shelf
(Table 1, Fig. 1) were sampled bimonthly for two years (Sept
1971 - July 1973). A Shipek benthic grab was used to obtain
five replicate samples at each station during each visit; each
grab sampled 0.04 m? of bottom surface area. Station
sediments were sieved, described, and classified to type dur-
ing 8 of the 12 sampling periods (Sept 1971; Jan, May, and
Sept 1972 excluded) (Gallagher, 1977). The substratum at all
stations was sand or sand-shell, lacking attached vegetation.
Station | was located in an area of high wave energy on the
beach terrace; sediments were fine to very fine, moderately
well-sorted, gray terrigenous sands (type 1), often very com-
pacted. Station II was offshore of Station | in a ‘‘trough’’ be-
tween the beach terrace and Pierce Shoal; sediments were
very coarse, poorly sorted sands containing very little mud
(type 2), sometimes with numerous large shell fragments;
mean particle size in sediment samples at Station II fluctuated
between those of Stations IV and V, indicating patchiness in
that area. Station III, located atop Pierce Shoal, was farthest
offshore (2 mi) but was the shallowest station; sediments were
medium-grained, moderately well-sorted calcareous sands
(type 3); large shell fragments, common in the trough, were
absent at Station Ill. Station IV was located in the trough

American Malacological Bulletin, No. 7(1) (1989):57-64

57

58 AMER. MALAC. BULL. 7(1) (1989)

Table 1. Coordinates and mean depths of benthic stations.

Mean depth
Station Coordinates* (m)
| 27921.3'N, 80°14.1.W 8.4
I 27°21.6’N, 80°13.2’W 11.2
ll 27922.0'N, 80°12.4’'W 71
IV 27920.7’N, 80°12.8’'W 10.9
V 27922.9'N, 80°13.9'W 10.8

*U.S.C.G.S. Chart No. 1247, dated 1969.

south of Station Il; sediments were very similar to those at
Station II (type 2), but sands were slightly less coarse; large
shell fragments also were present. Station V, another trough
station, was located north of Station II; sediments were very
coarse, poorly sorted, slightly muddy, calcareous sandy-shell
gravels (type 4).

Samples were preserved in 10% buffered formalin
when collected and then were washed through a 0.71 mm U.S.
Standard Sieve screen. Materials retained on the screen were
preserved in alcohol, sorted to higher taxonomic categories
using a binocular dissecting microscope, and transferred to
specialists for identification and enumeration. | examined all
mollusks from the samples and prepared a technical report
on species composition and abundance (Lyons, in press). The
specimens are deposited in the FDNR Marine Invertebrate
Collection at St. Petersburg, Florida.

Species were designated as characteristic of the study
site based upon occurrence during at least 6 of the 12 sampl-
ing periods. Interstation similarities of the fauna were deter-
mined using Czekanowski similarity coefficients (Clifford and
Stephenson, 1975) derived from raw bimonthly abundances
(Q mode) of the characteristic species at each station (five
replicate grabs pooled) during each of eight sampling periods
when sediments were analyzed. Station sediments likewise
were examined for similarity using percentages by weight of
particle sizes (@) of the samples during each of the same
eight periods. Resultant dendrograms of similarity based on
fauna and on sediments were compared for evidence of faunal
affinity for sediment types.

RESULTS

One hundred thirty-six molluscan species comprising
4135 living specimens were collected with the benthic grab.
Station I, which yielded only 40 specimens in 16 species dur-
ing 11 sampling periods (one atypical sample excluded), was
eliminated from further analyses. Station III, the shoal station,
yielded 697 specimens in 23 species. Greatest abundance
and diversity occurred at the trough stations: Station II—1139
specimens, 72 species; Station |V—841 specimens, 79
species; Station V—1418 specimens, 70 species. By class, 54
bivalve species contributed 75.5% of all specimens, followed
by 78 gastropods (18.4%), 3 polyplacophorans (5.2%), and
1 scaphopod (0.9%). Identities and abundances of all species
were reported elsewhere (Lyons, in press).

Only 33 of the 136 species (24%) were collected dur-

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oO.

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STUART AG

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6
KILOMETERS

Fig. 1. Location of offshore benthic sampling stations at Hutchinson
Island, central Florida east coast; depth contours in meters.

ing at least 6 of the 12 sampling periods, but those
characteristic species contributed 90% of all specimens ob-
tained during the study. The characteristic species, which in-
cluded 16 bivalves, 13 gastropods, 3 polyplacophorans, and
1 scaphopod, contributed greater proportions of the total fauna
at Stations Ill and V than at Stations Il and IV (Table 2).
Czekanowski similarity coefficients derived from abun-
dances of the characteristic species were used to construct
a dendrogram of relationships among samples collected at
Stations II-V during eight sampling periods (Fig. 2). Three
groups were evident: one group contained all of the Station
lll samples, one contained six Station Il samples and seven

LYONS: WESTERN ATLANTIC CRASS/NELLA 59

Table 2. Abundance and frequency of occurrence of characteristic molluscan species in grab samples, Hutchinson Island Stations II-V, Sept
1971 - July 1973 (n = number of specimens; m = number of months of occurrence).

Station

Species Total ll ll IV V

n m n m n m n m n m
Crassinella lunulata (Conrad, 1834) 1373 12 367 10 117 7 889 12
C. dupliniana (Dall, 1903) 580 12 55 10 458 12 51 10 16 6
Chione intapurpurea (Conrad, 1849) 350 12 137 12 12 7 107 10 94 9
Caecum cooperi S. Smith, 1860 189 12 69 11 37 8 44 9 39 10
Glycymeris spectralis Nicol, 1952 148 12 13 6 82 12 52 10 1 1
Ischnochiton niveus Ferreira, 1987 123 12 47 10 47 9 29 10
Chione grus (Holmes, 1858) 110 6 5 2 67 2 38 5
Calyptraea centralis (Conrad, 1841) 84 12 29 7 37 11 18 8
Caecum strigosum de Folin, 1868 82 11 44 5 18 7 20 8
Chaetopleura apiculata (Say, 1834) 72 12 17 7 31 5 24 9
Arene tricarinata (Stearns, 1872) 66 10 17 7 1 1 18 6 30 5
Macoma brevifrons (Say, 1834) 55 12 21 9 21 8 13 8
Ervilia concentrica (Holmes, 1860) 49 6 4 1 43 5 2 2
Pleuromeris tridentata (Say, 1826) 44 10 16 4 10 7 13 6 5 2
Corbula barrattiana C.B. Adams, 1852 41 11 6 3 16 7 19 9
Graptacme calamus (Dall, 1889) 39 9 3 3 22 9 14 3
Pteromeris perplana (Conrad, 1841) 37 12 10 6 1 1 25 11 1 1
Nucula proxima Say, 1822 31 10 7 3 2 2 22 8
Caecum floridanum Stimpson, 1851 25 10 12 8 13 6
Abra aequalis (Say, 1822) 24 6 7 3 4 2 13 5
Polygyreulima sp. A 22 10 12 8 7 4 3 2
Finella adamsi (Dall, 1889) 21 6 11 3 2 2 4 2 4 2
Ischnochiton hartmeyeri Thiele, 1910 20 16 8 3 9 5 3 2
Suturoglypta iontha (Ravenel, 1861) 19 7 5 4 3 2 11 5
Astyris lunata (Say, 1826) 16 8 3 2 4 1 9 6
Nassarius consensus (Ravenel, 1861) 14 7 6 3 2 2 6 5
Chama congregata (Conrad, 1833) 13 9 3 2 7 5 3 3
Semele bellastriata (Conrad, 1837) 12 8 7 5 2 2 3 2
Semelina nuculoides (Conrad, 1841) 10 6 2 2 6 4 2 1
Olivella floralia (Duclos, 1853) 9 7 1 1 3 2 5 5
Prunum roscidum (Redfield, 1860) 9 7 2 2 7 5
Tivela floridana Rehder, 1939 9 7 2 2 L 5
Seila cf. S. adamsi (H.C. Lea, 1845) 8 7 2 2 1 1 5 5
Total specimens (33 spp.) 3704 948 681 743 1332
Percent all specimens at station (128 spp.) 90.4 83.2 97.7 88.3 93.9

Station IV samples, and one contained all Station V samples
in addition to the two remaining Station II samples and one
Station IV sample. Similarity coefficients derived from
sediments produced a dendrogram with virtually the same
groupings (Fig. 2). The two Station Il samples that clustered
with those of Station V were the same two samples placed
there by species composition and abundance. Sediments of
the atypical Station IV sample (IV-2) did not cluster with Sta-
tion V sediments, but raw data values reveal that the sample
contained approximately 5% more large shell particles than
did other sediment samples at Station IV. That slight increase
evidently was sufficient to support a species group more
typically associated with sediments found at Station V.
The most abundant mollusks at Hutchinson Island
were two species of the bivalve genus Crassinella
(Crassatellidae). C. Junulata and C. dupliniana together con-
stituted 47% of all specimens; considering only the
characteristic species, that influence increased to 53%.

Crassinella lunulata was most abundant among the
large shell particles in sediments of Station V but also was
abundant occasionally at Stations Il and IV. Of 367 C. /unulata
recorded at Station II, 347 (95%) occurred in the two samples
in which sediments were of the type found at Station V (see
Fig. 2). At Station IV, 44 of 117 C. /unulata occurred in the sam-
ple with sediments that contained 5% more large shell par-
ticles, and 67 specimens occurred in one other sample.
Sediments of the latter sample were not analyzed, but the
associated fauna included large numbers of several other
mollusks (e.g. Chaetopleura apiculata, Chione grus) typically
associated with large shell particles. Thus, 95% of the C.
lunulata at Station IV also occurred in two samples with
sediments more similar to those at Station V. Together, the
12 Station V samples and the two samples each from Sta-
tions Il and IV contained 98% of all C. /unulata. Seventy-nine
percent of all C. dupliniana occurred among the well-sorted
medium sands of Station Ill, and the remainder of the spec-

60 AMER. MALAC. BULL. 7(1) (1989)

CHARACTERISTIC SPECIES SEDIMENTS
Similarity Level STATION Similarity Level
0 25 50 : 75) 100 100 75 50 25

—aaemmeroc tne (a

eral _— I-12 >) b

>———v-2 |

J
L

io a a © |I-8 “3
hms 3
Fig. 2. Dendrograms of Czekanowski similarity coefficients (Q mode)
based upon raw abundances of 33 characteristic species (left) and
percent size composition of sediments (right), showing cor-
respondence between species associations and sediments at four
stations (II-V) sampled bimonthly, Sept 1971-July 1973 (4 months
excluded).

imens were distributed among trough Stations Il, IV, and V.

To test the dependence of the two Crassinella species
on certain sediments, abundance data for each species were
examined by station and by sediment type using samples col-
lected during the eight periods when sediments were ana-
lyzed. As defined by Gallagher (1977), type 2 sediments oc-
curred at Stations II (Six samples) and IV (all samples), type
3 sediments occurred only at Station Ill, and type 4 sediments
occurred at Station V (all samples) and occasionally at Sta-
tion Il (two samples). For this analysis, however, the November
1971 sample (I\V-2) at Station IV was considered a type 4 sedi-
ment. Although sediments in that sample did not cluster with
those at Station V, the sample did contain greater quantities
of large shell particles, and the fauna clustered with fauna
typical of Station V (Fig. 2). Species abundance data were
equalized by converting to average catch per sample; each
station was sampled eight times, so abundances at Stations
Ill and V each were divided by 8, and combined abundances
at Stations Il and IV were divided by 16. Types 2, 3, and 4
sediments occurred in 13, 8, and 11 samples, respectively.

Relative abundances of the two species of Crassinella
in eight samples at each station differed little from those in
the total 12 samples (Table 3). However, average catch by sedi-
ment type demonstrated clearly the affinity of C. /unulata for
type 4 sediments (Table 4).

Information on recruitment, growth, and longevity was
discerned from size frequency distributions of the two
Crassinella species. Height of the largest specimen of C.
lunulata was 8.5 mm, but specimens seldom exceeded 6 mm
except during September 1971 (Fig. 3). Small (<1.5 mm)
juveniles occurred during all sampling periods except
September 1971, suggesting some year-round recruitment into

the population. Scarcity of small specimens in September
1971 samples is not understood. Such specimens must have
been present in the study area to produce the 2-4 mm
specimens common during following sampling periods.
Bimodality of sizes during September 1972 and May and July
1973 indicates successful ‘‘sets’’ prior to those months, pro-
bably during spring through fall 1972 and spring 1973. Only
a few individuals in September 1972 survived to sizes domi-
nant in September 1971, and the 1971-72 year class never at-
tained the maximum size of the September 1971 sample.
However, the 1972-73 year class was very successful, and the
larger size group of July 1973 probably would have attained
maximum size by fall 1973. Together, the data indicate a life
span of 12-18 months.

Height of the largest dead shell of Crassinella duplini-
ana was 3.4 mm, but no living specimens larger than 2.8 mm
were obtained (Fig. 4). Smallest specimens (1.0-1.2 mm) oc-
curred during May through November of each year, indicating
recruitment during warm months, and largest specimens
(2.6-2.8 mm) usually occurred in May and July. Although
cohort growth progressions among intermediate sizes
(1.3-2.5 mm) were less evident, C. dupliniana recruits seemed
to grow to 2.5-2.8 mm in about 12 months, and maximum size
(3.4 mm) probably was attained by specimens that survived
for a few more months.

DISCUSSION

Although the 136 species of mollusks collected in sand-
shell substrates off Hutchinson Island suggest a diverse fauna,
most species were relatively rare. Some species were scarce

PERCENT FREQUENCY/MONTH

HEIGHT (mm)

Fig. 3. Size frequency (0.3 mm increments shell height) of Crassinella
lunulata at Hutchinson Island, Sept 1971-July 1973.

|
|
|

LYONS: WESTERN ATLANTIC CRASS/NELLA 61

Table 3. Relative abundance (%) of Crassinella dupliniana and C. /unulata in 12 and 8 bimonthly samples at Hutchinson

Island benthic stations (catch combined for stations II and IV).

Total
Specimens % Occurrence by Station
(100%) Il, IV Wl Vv
Species 12 mo. 8 mo. 12 mo. 8mo. 12 mo. 8 mo. 12 mo. 8mo.
C. dupliniana 580 365 18.3 19.2 79.0 775 28 33
C. lunulata 1373 1006 35.3 40.7 0.0 0.0 64.7 59.3

representatives from nearby estuarine and deeper offshore
assemblages, and others were juveniles of tropical forms that
recruited to the area via the nearby Florida Current during
spring and summer but died during fall and winter. However,
most rare taxa seemed to be shallow shelf species typically
affiliated with hard substrata. Many of those species associate
with algae or sponges that exist, within the study area, prin-
cipally on the scarce and widely scattered shells of larger dead
mollusks (e.g. species of Mercenaria, Busycon, Pleuroploca,
Hexaplex, and Argopecten). Expectedly, the scattered hard-
substratum habitat was not sampled adequately with the grab.

Sediments at the study site are typical of those that
border the coast of east central Florida. The shallow shelf is
a submerged sedimentary plain, generally of low relief but
with ridge-like linear shoals resting on the seaward-dipping
platform (Meisburger and Duane, 1971). The linear shoals form
a small angle (most <35°) with the coast line and open
northward (Fig. 1). Such shoals usually are formed at the
shore-face in response to interactions between south-trending,
wind-driven surface currents and wave-generated bottom cur-
rents during winter storms. Offshore shoals represent previous
shore-face shoals detached by landward erosion (Duane et
al., 1972). Sediment types at the shoal and on the surround-
ing bottom in the study area essentially are the same as those
that occur throughout the inshore region between St. Lucie
Inlet and Cape Canaveral, Florida (Meisburger and Duane,
1971; Duane et a/., 1972).

The 33 characteristic species found in the coastal
oceanic environment at Hutchinson Island constitute a typical
molluscan assemblage of small forms adapted to sand and
shell-hash bottom sediments. However, differences in sedi-
ment composition strongly influenced the distributions of
species in that environment. The hard-packed fine to very fine
sands of the beach terrace supported a very sparse fauna,
but 8 of the 16 species collected there did not occur further
offshore. The consistently well-sorted medium sands of the
offshore shoal also supported relatively few species, but unlike
the beach terrace, several species were abundant there. The
bivalves Crassinella dupliniana, Glycymeris spectralis, Ervilia
concentrica, Semelina nuculoides, and Tivela floridana, and
the scaphopod Graptacme calamus, were much more abun-
dant at the shoal than elsewhere. Except for E. concentrica,
those species generally are scarce or absent in most Florida
sand-bottom assemblages but probably occur at other off-
shore shoals along east central Florida. Characteristic species
associated with large shell particles in the trough included
the bivalves C. /unulata, Nucula proxima, and Abra aequalis,
and several gastropods. Reasons for the occurrence of C.

ee N:25
Mar.
QO braves wore.
50
N: 86
May
ie) —
7 N:54
Jul.
es a
5

N: 118

PERCENT FREQUENCY / MONTH

HEIGHT (mm)

Fig. 4. Size frequency (0.1 mm increments shell height) of Crassinella
dupliniana at Hutchinson Island, Sept 1971-July 1973.

lunulata are discussed subsequently. N. proxima and A.
aequalis are deposit feeders that ingest fine particles trapped
among the very coarse sediments. Gastropods such as
Suturoglypta iontha, Astyris lunata, Prunum roscidum, and
Seila cf. S. adamsi utilize the coarse sediments for shelter and
feed upon other organisms associated with those sediments.
The more heterogeneous sediments in other trough samples
supported more species than did sediments with large shell
fragments. Species associated with those sediments includ-
ed a bivalve, Chione intapurpurea, and several sand-dwelling
gastropods, e.g. Caecum cooperi, C. floridanum, C. strigosum,
and Finella adamsi.

The dependence of species on particular sediments

62 AMER. MALAC. BULL. 7(1) (1989)

Table 4. Average (x N) and relative abundance (%) of Crassinella dupliniana and C.
lunulata in 8 bimonthly samples, by station and sediment type.

Station Sediment Type
Species Il, IV Wl V 2 3 4
x Specimens/Sample
C. dupliniana 44 35.4 1.5 3.5 35.4 3.3
C. lunulata 25.6 0.0 746 1.4 0.0 89.8
% Occurrence
C. dupliniana 19.2 775 3.3 12.6 775 9.9
C. lunulata 40.7 0.0 59.3 18 0.0 98.2

is exemplified by the two most abundant bivalves, Crassinella
dupliniana and C. lunulata. C. dupliniana was most abundant
among the well-sorted medium sands of the offshore shoal
and occurred less commonly in the trough. Although never
as dominant as at the shoal, medium sands always were com-
ponents of sediments at the trough stations and evidently oc-
curred there in quantities sufficient to support lesser numbers
of C. dupliniana. Conversely, C. /unulata did not occur at all
atop the shoal but was abundant in sediments with large shell
particles in the trough. Only one juvenile specimen of C.
lunulata and no C. dupliniana occurred on the compacted fine
to very fine sands of the beach terrace. Harry (1966)
documented the affinity of C. /unulata for sediments with a
high percentage of coarse shell particles. C. mactracea
(Linsley, 1845), a species very similar to C. /unulata in mor-
phology and maximum size, lives in gravel communities from
New York to Massachusetts (Allen, 1968). Both species use
a delicate byssus to attach to surface substratum (Harry, 1966;
Allen, 1968). Harry (1966) speculated that C. /unulata also may
burrow. However, specimens of C. /unulata in most en-
vironments are of similar size or slightly larger than the shell
fragments among which they live. Consequently, it seems
reasonable to propose that C. /unulata lives almost interstitially
among the large shell fragments, which might provide sup-
port, protection from predators, and camouflage. The smaller
species, C. dupliniana, might derive similar benefits among
sediments of correspondingly smaller size.

The requirement for settlement on certain sediments
poses a recruitment problem for larvae of many bivalves.
Crassinella dupliniana, especially, could have a problem
because its preferred sediment, well-sorted medium sands,
occurs principally on the relatively uncommon shoals that
border the coast. Larvae generally distributed in the eddy cir-
culation over the eastern Florida shelf might have little chance
of encountering those shoals. Some mollusks with special
habitat requirements solve the recruitment problem by ab-
breviating or eliminating the planktonic larval period. Harry
(1966) proposed that some Crassatellidae, e.g. C. Junulata and
Eucrassatella speciosa (A. Adams, 1852), brood eggs at least
during early development. Cuna dailli Vanatta, 1904, a species
sometimes placed in Crassatellidae (Moore, 1957) and
sometimes in Condylocardiidae (Abbott, 1974), has been
demonstrated to be ovoviviparous (Moore, 1961). Partial or
complete ovoviviparity that shortens or eliminates the
planktonic larval period of Crassinella species would assure
that young were released at or near appropriate sediments.

The fact that most small juvenile Crassinella at Hutchinson
Island occurred in the same sediments occupied by adults
supports that hypothesis.

Unfortunately, no evidence of broodings can be
obtained from the specimens of Crassinella used in this study.
The specimens were examined in 1975 and were not in-
spected for evidence of brooding then. The 1953 living
specimens of the two species were accompanied by 12,762
paired valves of dead specimens. Because most dead
specimens were sealed and appeared fresh, it was necessary
to dry and open the specimens to discern the incidence of
live-collected material; most tissues were damaged or
destroyed during that process.

Recruitment of both Crassinella species occurred at
Hutchinson Island during much of the year. Egg-bearing C.
lunulata have been reported off eastern Florida during
September and off Texas during October (Harry, 1966). A
September spawn supports the fall recruitment of C. /unulata
suggested by some size frequency data from Hutchinson
Island, but earlier spawns during other warm months by C.
lunulata and by C. dupliniana also are indicated.

Crassinella dupliniana increases to five the number of
Recent species recognized in the northwestern Atlantic
Ocean. Harry (1966) recognized only two Recent species of
northwestern Atlantic Crassinella, C. lunulata and C. mar-
tinicensis (d’Orbigny, 1846); Abbott (1974) followed Harry’s
classification. However, Allen (1968) maintained that the
southern C. /unulata and the northern C. mactracea are
separate species, although he did not mention characters
which distinguish them. Because the status of those two taxa
is uncertain, | maintain them separately here. C. /unulata oc-
curs either from Massachusetts (Abbott, 1974) or from North
Carolina (Allen, 1968) to Florida, Texas, Bermuda, and Brazil.
Most recently, Coan (1984) recognized C. aduncata Weisbord,
1964, originally described as a Pliocene fossil, among the liv-
ing fauna of the Caribbean Sea.

Because Crassinella dupliniana has not been reported
in literature on Recent mollusks, it is illustrated here and com-
pared with congeners. In a paper submitted in 1975, Ward
and Blackwelder (1987) stated that the type specimen of C.
dupliniana might be lost, but | examined syntypes (USNM
114922) at the National Museum of Natural History which are
identical to the Hutchinson Island specimens. Although
dissimilar in outline to most other species of the genus, C.
dupliniana shares with them the external cellular texture,
unique to shells of Crassinella, described by Harry (1966) and

LYONS: WESTERN ATLANTIC CRASS/NELLA 63

Figs. 5-7. Left (upper) and right (lower) valves of three species of Crassinella from Florida. 5. C. dupliniana, height 2.8 mm, Hutchinson Island
Station IV, July 1972. 6. C. /unulata, height 2.7 mm, Hutchinson Island Station Il, January 1972. 7. C. martinicensis, height 2.7 mm, off Clear-
water, Florida, depth 44 m. All specimens deposited in FDNR Marine Invertebrate Collection.

Coan (1979). C. dupliniana (Fig. 5) is much smaller and has
amore acute apical angle than C. /unulata and C. mactracea.
The height of Dall’s largest specimen of C. dupliniana was
3.2 mm and that of the largest specimen from Hutchinson
Island was 3.4 mm, whereas maximum heights of C. /unu/ata
and C. mactracea approach 10 mm (Allen, 1968). Valves of
C. lunulata (Fig. 6) are quite compressed and have about 3
broad concentric ridges per millimeter of shell height, whereas
valves of C. dupliniana are more swollen, and concentric
ridges are finer and more closely spaced, averaging 6-7 per
millimeter. C. martinicensis (Fig. 7), a species that occurs off
both Florida coasts in 20-80 m depths, resembles the broad-
ly triangular C. /unulata and C. mactracea more than it does
the acute C. dupliniana. Harry (1966) reported a maximum
height of 2.7 mm for C. martinicensis, but Florida specimens
attain a maximum height of about 3.0 mm. Valves of C. mar-
tinicensis have 4-5 concentric ridges per millimeter of height;
the ridges often are spaced irregularly, and several secon-
dary ridges sometimes occur between them.

Crassinella dupliniana’ most resembles certain
specimens of C. nuculiformis Berry, 1940, a narrowly ovate,
inflated species that occurs from Baja California to Ecuador
(Coan, 1979). The 2.9 mm valve of C. nuculiformis illustrated
by Coan (1979: fig. 10) is strikingly similar to those of C. dupli-

niana. Coan (1979) noted that C. nuculiformis attains a height
of 6.4 mm, but specimens from the upper Gulf of California
(including the specimen in his fig. 10) were smaller, general-
ly <3 mm. According to Coan (1984), C. nuculiformis is very
similar to C. maldonadoensis (Pilsbry, 1897) from Uruguay to
Argentina in the southwestern Atlantic, but the latter species
has less prominent umbones and its concentric ribs fade more
quickly toward the ventral margin. C. adamsi Olsson, 1961,
an eastern Pacific species that attains a height of 3.6 mm,
also is ovate but is proportionally longer than C. nuculiformis
and C. dupliniana. Olsson (1961) mentioned an undescribed
species similar to C. adamsi from the Caribbean coast of
Panama. That species probably is C. aduncata, which differs
from C. adamsi ‘‘in attaining a larger size, having a more
abrupt posterior slope, and . . . more prominent concentric
ribs’ (Coan, 1984: 165).

In addition to the Hutchinson Island material, | have
examined specimens of Crassinella dupliniana from off Cape
Canaveral, Florida, from the Gulf of Mexico off western Florida
(both FDNR Marine Invertebrate Collection), and from beach
drift at Hunting Island State Park, South Carolina. These
records indicate that C. dupliniana probably lives among ap-
propriate sediments throughout much of the warm-temperate
Carolinian Province.

64 AMER. MALAC. BULL. 7(1) (1989)

Information on sediment associations of the two
Crassinella species could prove useful for interpreting
paleoenvironments. C. /unulata (Conrad, 1834), originally
described as a fossil from the Pliocene Yorktown Formation
of Suffolk, Virginia, occurs extensively in Pliocene and
Pleistocene deposits of the southeastern United States (Gard-
ner, 1944). C. dupliniana (Dall, 1903), originally described from
the Pliocene Duplin Formation (now considered Yorktown) of
Natural Well, Duplin County, North Carolina, also has been
reported from Pliocene and early Pleistocene deposits in
Virginia, the Carolinas, and north and south Florida (Dall,
1903; Mansfield, 1931, 1932; Gardner, 1944; Dubar and Taylor,
1962; Stanley, 1986; Ward and Blackwelder, 1987). High in-
cidence of either species in Neogene strata probably would
indicate environments similar to those with which they
associated at Hutchinson Island.

ACKNOWLEDGMENTS

Robert M. Gallagher and colleagues, Applied Biology, Inc.,
Jensen Beach, Florida, collected the samples, analyzed the
sediments, and transferred the mollusks to me for study. David K.
Camp, Walter C. Jaap, and Gayle Plaia, FDNR, assisted with com-
puter analyses of similarity. Dr. Earnest Truby, FDNR, made the SEM
photographs. Dr. Thomas R. Waller allowed me to examine specimens
of Crassinella dupliniana at the U. S. National Museum of Natural
History, and Dr. Lyle C. Campbell, University of South Carolina,
Spartanburg, provided Recent specimens from South Carolina. The
study was funded in part by Florida Power and Light Company, Inc.
All are gratefully thanked.

LITERATURE CITED

Abbott, R. T. 1974. American Seashells, 2nd ed. Van Nostrand
Reinhold Company, New York. 663 pp.

Allen, J. A. 1968. The functional morphology of Crassinella mactracea
(Linsley) (Bivalvia: Astartacea). Proceedings of the
Malacological Society of London 38:27-40.

Clifford, H. T. and W. Stephenson. 1975. An Introduction to Numerical
Classification. Academic Press, New York. 229 pp.

Coan, E. V. 1979. Recent eastern Pacific species of the crassatellid
bivalve genus Crassinella. Veliger 22(1):1-11.

Coan, E. V. 1984. The Recent Crassatellinae of the eastern Pacific,
with some notes on Crassinella. Veliger 26(3):153-169.
Conrad, T. A. 1834. Description of new Tertiary fossils from the
southern states. Journal of the Academy of Natural Sciences

of Philadelphia, series 1, 7:130-178.

Dall, W. H. 1903. Contributions to the Tertiary fauna of Florida with
especial reference to the silex beds of Tampa and the Pliocene
beds of the Caloosahatchie River. Transactions of the Wagner

Free Institute of Science of Philadelphia 3(6):1219-1654.

Duane, D. B., M. E. Field, E. P Meisburger, D. J. Swift and S. J.
Williams. 1972. Linear shoals on the Atlantic inner continen-
tal shelf, Florida to Long Island. In: Shelf Sediment Transport,
D. J. Swift, D. B. Duane, and O. H. Pilkey, eds. pp. 447-498.
Dowden, Hutchinson and Ross, Inc., Stroudsburg,
Pennsylvania.

Dubar, J. R. and D. S. Taylor. 1962. Paleoecology of the Choctaw-
hatchee deposits, Jackson Bluff, Florida. Transactions of the
Gulf Coast Association of Geological Societies 12:349-376.

Gallagher, R. M. 1977. Nearshore marine ecology at Hutchinson
Island, Florida: 1971-1974. Il. Sediments. Florida Marine
Research Publications 23:6-24.

Gardner, J. 1944. Mollusca from the Miocene and lower Pliocene of
Virginia and North Carolina. Part 1. Pelecypoda. United States
Geological Survey Professional Paper 199-A:1-178.

Harry, H. W. 1966. Studies on bivalve molluscs of the genus
Crassinella in the north-western Gulf of Mexico: anatomy,
ecology and systematics. Publications of the Institute of Marine
Science of the University of Texas 11:65-89.

Lyons, W. G. In press. Nearshore marine ecology at Hutchinson
Island, Florida: 1971-1974. XI. Mollusks. Florida Marine Research
Publications.

Mansfield, W. C. 1931. Some Tertiary mollusks from south Florida.
Proceedings of the United States National Museum 79(21):1-12,
pls. 1-4.

Mansfield, W. C. 1932. Miocene pelecypods of the Choctawhatchee
Formation of Florida. Florida Geological Survey Geological
Bulletin 8:1-240.

McCloskey, L. R. 1970. The dynamics of the community associated
with a marine scleractinian coral. Internationale Revue der
Gesamten Hydrobiologie 55(1):13-81.

Meisburger, E. P. and D. B. Duane. 1971. Geomorphology and
sediments of the inner continental shelf, Palm Beach to Cape
Kennedy, Florida. United States Army Corps of Engineers
Technical Memorandum 34:1-111.

Moore, D. R. 1957. A note on Cuna dalli. Nautilus 70(4):123-125.
Moore, D. R. 1961. The marine and brackish water Mollusca of the
State of Mississippi. Gu/f Research Reports 1(1):1-58.
Reed, J. K. and P. M. Mikkelsen. 1987. The molluscan community
associated with the scleractinian coral Oculina varicosa.

Bulletin of Marine Science 40(1):99-131.

Stanley, S. M. 1986. Anatomy of a regional mass extinction: Plio-
Pleistocene decimation of the western Atlantic bivalve fauna.
PALAIOS 1:17-36.

Ward, L. W. and B. W. Blackwelder. 1987. Late Pliocene and early
Pleistocene Mollusca from the James City and Chowan River
Formations at the Lee Creek Mine. In: Geology and Paleon-
tology of the Lee Creek Mine, North Carolina, II. C. E. Ray, ed.
pp. 113-283. Smithsonian Contributions to Paleobiology 61.

Date of manuscript acceptance: 23 August 1988

TEMPORAL VARIATION IN MICROSTRUCTURE OF THE
INNER SHELL SURFACE OF CORBICULA FLUMINEA
(BIVALVIA: HETERODONTA)

ANTONIETO TAN TIU
HARBOR BRANCH OCEANOGRAPHIC INSTITUTION
DIVISION OF APPLIED BIOLOGY
5600 OLD DIXIE HIGHWAY
FT. PIERCE, FLORIDA 34946, U.S.A.

ROBERT S. PREZANT
DEPARTMENT OF BIOLOGY
INDIANA UNIVERSITY OF PENNSYLVANIA
INDIANA, PENNSYLVANIA 15705, U.S.A.

ABSTRACT

Temporal variation of shell microstructure with emphasis on the inner shell surface was examined
in caged and noncaged Corbicula fluminea (Muller) from the Leaf River, Mississippi. Shell structure
in the outer shell layer, overlain by the periostracum, exhibited distinct seasonal variation from crossed-
lamellar in warmer months to structures resembling cone complex crossed-lamellar in cooler months.
Other variations associated with season were subtle, involving only the inner shell surface microstruc-
tures, such as replacement of well developed lamellae in warmer months by pitted, deformed or reticulate
microstructures in cooler months. The shell microstructure ventral to the pallial line is of possible use
in taxonomic and phylogenetic analyses of the Corbiculacea and could also be of value as an en-
vironmental monitor because the microstructures in this region were less variable than those dorsal

to the pallial line.

Previous workers [i.e. Carter (1980)] have hypothesized
that major variations in shell microstructures among bivalves
are largely biologically controlled (i.e. more or less indepen-
dent of environmental influences) and have developed adap-
tive edges towards resistance to abrasion or fracture, energy
economies of secretion, etc. There are however exceptions
to this generalization. Lutz and Rhoads (1979, 1980) and Lutz
and Clark (1984) demonstrated that the microstructure of the
inner shell layer of Geukensia demissa (Dillwyn) varied with
season and latitude. Moreover, Tan Tiu and Prezant (1987)
have shown that variation in the microstructure of the inner
shell surface of G. od. granosissima (Sowerby) occurred within
a small geographical area. Likewise, ultrastructure of the in-
ner shell surface in Polymesoda caroliniana (Bosc) can reflect
seasonal and/or habitat variation (Tan Tiu, 1987, 1988). Thus,
environmental variation can directly or indirectly influence the
deposition of shell microstructural components resulting in
at least surficial modifications.

A brief review of bivalve shell microstructural studies

will reveal the phylogenetic and ecological importance of such
structural variations. The information derived from en-
vironmental modification of bivalve shell microstructure can
help in understanding not only molluscan phylogeny (Carter,
1980), but also the historical (perhaps paleontological) events
that brought about these changes (Lutz and Rhoads, 1980;
Rhoads and Lutz, 1980). Realization of the full potential of
shell microstructure patterns as a taxonomic tool and as a
recorder of environmental change, depends on examination
of bivalve shell microstructural variations (Carter, 1980). This
paper reports temporal variations in the microstructure of the
inner shell surface of caged and noncaged Corbicula fluminea
(Muller) in the Leaf River, Belleville, Perry County, Mississippi,
U.S.A.

MATERIALS AND METHODS

Caged and noncaged specimens of Corbicula fluminea
were sampled seasonally from the south bank of the Leaf

American Malacological Bulletin, Vol. 7(1) (1989):65-71

65

66 AMER. MALAC. BULL. 7(1) (1989)

River (Belleville, Perry County, Mississippi) from June 1985
to June 1986. Additional noncaged samples were collected
in October 1985 and January 1986. Dates of collection and
number and lengths of specimens examined are presented
in Table 1.

The source of caged samples was a ‘‘natural’’ popula-
tion of Corbicula fluminea collected in June 1985 from the
same site in the Leaf River. Forty-five marked clams were
placed in each of eight cylindrical wire cages made of
galvanized iron (30 cm long, 20 cm diameter, 0.7 cm mesh)
and returned to the original collecting site in the Leaf River.
Each cage was fastened to an iron pole using wires, and
cages were set about two meters apart. Each pole was forced
into the substratum, the bottom of the attached cage touching
or slightly below the surface of the substratum.

Clams from each of two cages were shucked in the
field at seasonal intervals (Table 1) and the shells were fixed
separately in absolute ethanol. Select fractured and unfrac-
tured shell samples were dried in a Denton DCP-1 critical point
drier using liquid CO». as a transfer agent, mounted on
aluminum stubs using silver paint, coated with gold in a
Polaron SEM Coating Unit E5100, and examined at 30 KV us-
ing an AMR 1000A scanning electron microscope. Nine areas
of the inner shell surface were examined and compared, from
the ventral shell margin to the umbo (Fig. 1). Whenever possi-
ble, terminology of microstructures of inner shell surfaces cor-
related with that proposed by Carter and Clark (1985).

Several biological and environmental variables were
measured (Tan Tiu, 1987) but only monthly temperature of bot-
tom water (+ 1°C) in the Leaf River is presented here.

RESULTS

A. SHELL MICROSTRUCTURE

Microstructure of the inner shell surface varied distinct-
ly from the ventrum (Area A) to the dorsum (Area |) (Figs. 2
- 10), but not along the curved anterior posterior axis. Distinct
as well as subtle seasonal variations in the microstructure of

Table 1. Lengths (mm) of Corbicula fluminea from Leaf River, whose
shell microstructures were examined by scanning electron
microscopy.

Date Mean + 1 Standard Range Total Clams
Deviation Examined
Caged
Sept 85 2534+ 3.1 15.8 - 285 30
Dec 85 277 + 3.2 22.7 - 358 29
March 86 304+ 28 26.1 - 37.9 30
June 86 30.3 + 09 29.0 - 31.6 10
Noncaged
June 85 219+ 2.2 18.0 - 248 10
Sept 85 1914+ 45 10.3 - 23.7 10
Oct 85 24.2 + 10.0 9.5 - 38.0 28
Dec 85 238 + 76 12.5 - 376 10
Jan 86 250+ 73 10.1 - 38.0 10
March 86 294+ 45 21.5 - 38.0 9
June 86 319 + 19 278 - 34.0 10

rPOooumMmnegat —

Fig. 1. Right valve of Corbicula fluminea. Areas of the shell surface
examined are marked by dots, corresponding to the letters on the
right: A, internal surface area overlain by periostracum; B, area just
dorsal to Area A; C, area between Area B and pallial line; D, E, and
F, the ‘‘transition zone’; G, area at the level of ventral margin of ad-
ductor scars; H, area at the level of dorsal margin of adductor scars;
|, area near umbo.

the inner surfaces of shells are summarized in Table 2. With
a minor exception presented below, there were no differences
in the overall appearance and frequency of occurrence of
microstructures of the inner shell surface in caged and non-
caged Corbicula fluminea (Table 2).

1. OUTER SHELL LAYER

a. Area Overlain by Periostracum (Area A). Microstruc-
ture of the inner shell surface in the area overlain by the
periostracum can be divided into Crossed-Lamella One and
Microstructure C. Tertiary lamellae are apparently not
organized into broad secondary lamellae in Crossed-Lamella
One (Fig. 2). The first order lamellae in Crossed-Lamella One
are less than 40 »m wide, and are arranged diffusely. The
shell structure of Crossed-Lamella One approaches the
medium diffuse crossed lamellar structure of Carter and Clark
(1985). Microstructure C is a term of convenience that refers
collectively to spiral, pseudospiral or rosette arrangements
of laths surficially composing the crossed lamellar shell struc-
ture in area A (Fig. 11). The secondary lamellae on the deposi-
tional shell surface are arranged continuously into a spiral
in spiral crossed-lamellar structure, discontinuously or into op-
posing hemispheres (arcs) in pseudospiral crossed-lamellar
structure, and into irregularly arranged curved lamellae in
rosette crossed-lamellar structure. Spiral and pseudospiral
crossed-lamellar structures have been previously described
by Prezant and Tan Tiu (1986), and both structures approach
the cone complex crossed lamellar structure of Carter and
Clark (1985). While Crossed-Lamella One occurs throughout
the year, Microstructure C is present only in cooler months
(Oct to March 1986) (Table 2).

b. Areas Between Area A and Pallial Line (Areas B and
C). Laths in area B are arranged more or less regularly into

TAN TIU AND PREZANT: SHELL MICROSTRUCTURE OF CORBICULA 67

bee Cenblion Wi)

Fig. 2. Area A, overlain by periostracum, consisting of laths not organized into second order lamellae. Referred to as Crossed-Lamella One
in the text [horizontal field width (HFW) = 14 um]. Fig. 3. Area B, just dorsal to Area A consisting of secondary lamellae of opposing directions,
together with Fig. 4 (Area C) make up Crossed-Lamella Two referred to in the text (HFW = 14 um). Fig. 4. Area C, between Areas B and
pallial line with overlying organic matrix (HFW = 14 um). Fig. 5. Area D, just dorsal to pallial line (part of transition zone) consisting of narrow
lamellae (HFW = 14 um). Fig. 6. Area E, middle third of transition zone consisting of wide irregular lamellae (HFW = 14 um). Fig. 7. Area
F, just dorsal to Area E consisting of laths superimposed on irregularly fused lamellae (HFW = 14 um). Fig. 8. Area G, at level of ventral
margin of adductor scar consisting of irregularly fused lamellae that are perpendicular to shell surface. Referred to in the text as Complex
Crossed-Lamella One (HFW = 14 um). Fig. 9. Area H, at level of dorsal margin of adductor scar consists of overlapping broad lamellae, together
with Fig. 10 (Area |) below is referred to as Complex Crossed-Lamella Two (HFW = 14 um). Fig. 10. Area 1, near umbo (where tubules are
located) can be eroded (HFW = 14 um).

68 AMER. MALAC. BULL. 7(1) (1989)

second order lamellae, the latter in turn is arranged regular-
ly to form first order lamellae less than 10 um wide. Direction
of the second order lamellae is opposite that of adjacent first
order lamellae (Fig. 3). The shell structure in area B ap-
proaches the compressed crossed lamellar structure of Carter
and Clark (1985). The laths in area C are irregularly arranged
and apparently not organized into wide secondary lamellae
as in Crossed-Lamella One. The boundaries of the first order
lamellae are indistinct. Area C is often covered by an organic
sheet that renders the underlying structures indistinct, except
for the lamellar tips (Fig. 4). Microstructures of the inner shell
surfaces in both areas A and B are referred to as Crossed-
Lamella Two (Table 2). Loose or dense networks, which can
be granulated, are distributed evenly or patchily in areas B
and C. These networks are part of a continuum that is here
referred to as Reticulate Microstructures (Fig. 12, Table 2).
Frequency of occurrence of Reticulate Microstructures in
areas B and C is variable (Table 2).

2. INNER SHELL LAYER

a. Areas of the Transition Zone (Areas D, E and F).
These areas make up the transition zone, where the newly
deposited laths, destined to become the inner complex
crossed-lamella, are first formed over the pallial line. Incipient
as well as narrow and thin laths are observed in area D adja-
cent to the pallial myostracum (Fig. 5). Laths that are broad
and thick are often fused into irregular lamellae in Area E
(Fig. 6). In area F, the lamellae are broader, with incipient laths
over them (Fig. 7). The boundaries of first order lamellae on
the depositional surface of the transition zone are indistinct.
The structures composing the transition zone can also be
overlain with loosely arranged Reticulate Microstructures (Fig.
12). The latter is often very dense and obscures the underly-
ing original structures. Lamellae of the transition zone are
generally perforated, deformed (appearance similar to Fig.
13) or absent in December and March. Reticulate Microstruc-

tures in areas D - F did not show distinct seasonal variation.
However, as in areas B and C, the frequency of occurrence
of this microstructure increased when the frequency of oc-
currence of ‘‘well formed’’ structures (Figs. 5 - 7) in these
areas decreased.

b. Areas Dorsal to the Transition Zone (Areas G, H and
1). The shell structure in Area G, H and | is inconsistent, ap-
proaching either the irregular or cone complex crossed
lamellar structures of Carter and Clark (1985). Appearance
of the exposed lamellae of the inner surface of shell dorsal
to the pallial line is also variable. Among the commonly
observed microstructures, three are described here and con-
veniently named as Complex Crossed-Lamella One, Complex
Crossed-Lamella Two, and again Reticulate Microstructure.

Many of the lamellae are nearly perpendicular to the
inner surface of shell and are irregularly arranged in Complex-
Crossed Lamella One (Fig. 8). This microstructure is present
in June and absent in December in both caged and noncaged
clams.

The exposed ends of the lamellae in Complex Crossed-
Lamella Two are wide and broad, arranged such that they
overlap, one on top of the other like shingles (Fig. 9); they
also can be slightly eroded (Fig. 10). Seasonal frequency of
occurrence of this microstructure was lower in caged than
noncaged clams. When present in December and March, this
microstructure was eroded and deformed.

The appearance of Reticulate Microstructure in Areas
G to | varied. A loosely to densely packed thin stranded net-
work with or without granulations was present in varying
amounts throughout the year. In December and March
samples, this network had thicker strands with few granula-
tions (Fig. 12).

Tubules that penetrated the calcareous shell compo-
nent were consistently observed in the early dissoconch shell.
The shell tubules are filled with mantle extensions. The lat-
ter are at least occasionally bifurcate toward the shell exterior

Table 2. Temporal variation in microstructure of inner shell surface of Corbicula fluminea from Leaf River, Mississippi. Frequency
of occurrence expressed in classes where 0 = 0, 1 = 1 to 20, 2 = 21 to 40, 3 = 41 to 60, 4 = 61 to 80 and 5 = 81 to 100%.
C = Microstructure C, a collective new term for spirals, pseudospirals and rosettes; CL = Crossed-Lamella; RET = Reticulate
Microstructure; TRP = Transition Zone Lamellae are present; TRA = Transition Zone Lamellae are absent; CCL = Complex

Crossed-Lamella.

Area A Areas B-C Areas D-F Areas G-l
Cc CLI CL2 RET TRP TRA RET CCL1 CCL2 RET

Noncaged

June 85 0 5 5 0 5 0 0 2 2 2

Sept 85 0 5 5 0 5 0 0 0 4 1

Oct 85 1 5 5 0 5 0 2 0 3 2

Dec 85 3 2 5 1 3 3 0 0 3 3

Jan 86 3 2 5 0 3 1 2 0 0 4

March 86 2 3 5 1 4 0 2 1 0 4

June 86 0 5 5 1 5 0 2 1 1 3
Caged

Sept 85 0 5 5 0 5 0 1 2 1 2

Dec 85 3 3 5 1 3 1 1 0 2 3

March 86 3 3 4 2 3 0 3 0 1 5

June 86 0 5 4 2 4 0 4 2 0 4

TAN TIU AND PREZANT: SHELL MICROSTRUCTURE OF CORBICULA 69

FRA ANS
. ANS
Fig. 11. Spiral (S), pseudospiral (P) and rosette (R) microstructures

are present in Area A in both caged and noncaged clams during
cooler months (HFW = 23 um).

(Tan Tiu, 1987). Details of the microstructure and function of
the tubules are reported in a separate paper (Tan Tiu and
Prezant, 1988).

Some microstructures of the inner shell surface had
low frequency of occurrences. An example of a microstruc-
tural variant whose frequency of occurrence was low (less than
20% of total samples) and did not show distinct seasonal
variation is shown in figure 14. Lamellae in figure 14 are ar-
ranged in such a way that they resemble a pinwheel. Other
examples of such variants are described by Tan Tiu (1987).

B. WATER TEMPERATURE

The temperature of the bottom water in the Leaf River
was highest in August and lowest in January (Fig. 15). This
was the only environmental variable that showed a distinct
cyclic pattern.

DISCUSSION

The frequency of occurrence of Microstructure C in the
outer shell layer and ‘‘well formed”’ lamellae in the transition
zone of the inner shell layer of both caged and noncaged Cor-
bicula fluminea exhibited seasonal variation. The time of oc-
currence of these microstructures; however, were different.
Frequency of occurrence of Microstructure C was inversely
associated with bottom water temperature, while the presence
of ‘‘well formed’ lamellae in the transition zone was positively
associated with bottom water temperature.

Reticulate Microstructure observed in Corbicula
fluminea is similar in appearance to that observed in
Polymesoda caroliniana by Tan Tiu (1987, 1988). In both
species, an increase in occurrence of Reticulate Microstruc-
ture corresponded with a decrease in occurrence of other
microstructural ‘‘types’’ in the areas involved. Several studies
have suggested that valve closure results in calcium reab-
sorption from the inner shell surface (Crenshaw and Neff,

Fig. 12. Ultrastructure referred to as Reticulate Microstructure in the
text (HFW = 21 um). Fig. 13. Highly eroded lamellae predominating
in cooler months in Areas G to | (HFW = 19 um). Fig. 14. Pinwheel
arrangement of laths (HFW = 21 um).

1969; Lutz and Rhoads, 1979; Akberali and Trueman, 1985).
During valve closure under stressful or normal conditions,
bivalves shift to an anaerobic metabolic pathway to generate
ATP (Hochachka, 1980). The resulting acidic by-products,

70 AMER. MALAC. BULL. 7(1) (1989)

such as succinate, propionate, etc., of this metabolic pathway
are then buffered by carbonates in the calcareous shell
resulting in shell dissolution (Akberali and Trueman, 1985).
The Reticulate Microstructure with numerous spaces between
structures, could be a result of dissolution of the inner sur-
face of the shell consequent to valve closure as observed by
Prezant et a/. (1988) and as suggested by Akberali and
Trueman (1985).

While seasonal variation in appearance of microstruc-
tures dorsal to the pallial line consists mainly of “‘well formed’’
lamellae in warmer months being replaced by deformed
lamellae in cooler months, those microstructures ventral to
the pallial line in area A are ‘‘well formed’’ throughout all
seasons with Microstructure C predominating in cooler
months. This suggests that Crossed-Lamella One and
Microstructure C are inducible microstructures dependent on
varying environmental conditions associated with change in
season. Temperature of bottom water in Leaf River, which has
a distinct seasonal cycle, could play a major role in this
microstructural induction. The presence of ‘‘well formed”’
microstructures in all seasons ventral to the pallial line, but
only in warmer months dorsal to the pallial line, suggests that
shell growth is continuous along the shell margin, and periodic
on the inner shell layer dorsal to the pallial line. This was ex-
pected since winter in Mississippi is relatively short, and water
temperature in the Leaf River dropped below 10°C for only
a short time. According to Fritz and Lutz (1986), shell growth
of Corbicula fluminea in New Jersey occurs at water
temperatures above 10°C.

Prezant and Tan Tiu (1986) discovered spiral crossed-
lamellar microstructure in Summer and winter samples of Cor-
bicula fluminea from Strong River, Mississippi and
pseudospiral microstructure in a winter sample of C. fluminea
from Leaf River, Mississippi. These authors suggested a
seasonality of occurrence of pseudospiral microstructures in
Leaf River. Data presented here clearly indicate, for the first
time, that spiral crossed-lamellar microstructures (Fig. 11) are

45 +

35 +

25

°

©: (2. Gt 2 oe SS mS

MONTHS

Fig. 15. Monthly variation in temperature of the bottom water of Leaf
River at the collection site. Dots represent monthly records of water
temperature at sampling time, while vertical lines through the dots
represent monthly temperature ranges as measured by a maximum-
minimum thermometer.

found in C. fluminea in the Leaf River. Furthermore, spiral and
pseudospiral crossed-lamellar microstructures were deposited
only in cooler months by C. fluminea in the Leaf River. Similari-
ty of occurrence of these microstructures in caged and non-
caged Leaf River samples nullify the idea of Prezant and Tan
Tiu (1986) that Leaf River samples possessing pseudospiral
microstructure ‘‘drifted’’ per se from upstream habitats. Spiral
crossed-lamellar microstructure does not occur in the close-
ly related Polymesoda caroliniana or other corbiculids from
other habitats (Prezant and Tan Tiu, 1986; Tan Tiu, 1987, 1988).
Samples of P caroliniana collected in cooler months, however,
do exhibit a pseudospiral microstructure (Tan Tiu, 1987, 1988).
Polymesoda caroliniana is the only other corbiculid with data
on seasonal variation of shell microstructure (Tan Tiu, 1988).
Data on C. fluminea and P. caroliniana suggest that although
the microstructure at the internal shell margin overlain by
periostracum exhibits seasonal variation, it has taxonomic and
phylogenetic implications that warrant further consideration.

Dorsal to the pallial line, ‘‘well formed’’ structures
predominating in warmer months are usually replaced by
deformed or pitted structures in cooler months. This is similar
to phenomena observed in other bivalves. Wada (1960)
demonstrated that the size and shape of nacreous tablets
composing the inner shell layer of the bivalve Pinctada
martensii (Dunker) varied with season, being large and hex-
agonal in summer, and small, deformed and pitted during
winter. Lutz and Clark (1984) showed that nacreous tablets
of the inner shell layer of the Atlantic ribbed mussel Geukensia
demissa varied not only with season but also with latitude.
Further, Tan Tiu and Prezant (1987) showed that distinct dif-
ferences in the shape and size of nacreous tablets along the
inner surface of the inner shell layer of G. d. granosissima
can occur even within a small geographical area. In
Polymesoda caroliniana, Tan Tiu (1987, 1988) observed
seasonal and spatial variations of microstructures in the in-
ner surfaces of their shells. Because of high variability of the
microstructure dorsal to the pallial line brought about by
various factors, shell microstructure in this area has less tax-
onomic value than shell microstructure distal to the pallial
line. The significance of the pinwheel arrangement of lamellae
and other microstructural variations dorsal to the pallial line
(Tan Tiu, 1987) is currently unknown.

The guide to shell structure proposed by Carter and
Clark (1985) is clearly helpful in classifying shell structures
and inferring phylogeny. However, in Polymesoda caroliniana
(Tan Tiu, 1988) and Corbicula fluminea, where intrinsic or ex-
trinsic intraspecific variability in shell structure occurs, the use
of specific guides correlating specific taxa with specific shell
microstructures could be misleading.

ACKNOWLEDGMENTS

We thank Prof. Melbourne Carriker, Dr. Clement L. Counts,
Ill and the two anonymous critics for the review of the previous edi-
tions of our manuscript; Mr. Sheau-Yu Shu, Mr. Kashane Chalerm-
wat, Mr. Noel D’mello, Mr. Thomas Rogge, Miss Alene Minchew, Dr.
Kumar Prasanna for help in some sample collection; Dr. Raymond
Scheetz for help with scanning electron microscopy; Mr. and Mrs.

TAN TIU AND PREZANT: SHELL MICROSTRUCTURE OF CORBICULA 71

Franklin McRee and Miss Patricia McRee for allowing us to collect
samples and permitting us to place caged samples on their proper-
ty. This study was supported by a National Capital Shell Club Scholar-
ship, a grant from Sigma Xi and aid from the Department of Biological
Sciences, University of Southern Mississippi, U.S.A., and Research
and Faculty Development from the University of San Carlos, Philip-
pines granted to A. T. T. Publication costs have been funded by Har-
bor Branch Oceanographic Institution, Inc. (contribution no. 663).

LITERATURE CITED

Akberali, H. B. and E. R. Trueman. 1985. Effect of environmental stress
on marine bivalve molluscs. In: Advances in Marine Biology,
K. Wilbur, ed. pp. 101-198. Academic Press, London.

Carter, J. G. 1980. Environmental and biological controls of bivalve
shell mineralogy and microstructure. /n: Skeletal Growth of
Aquatic Organisms, D. C. Rhoads and R. A. Lutz, eds. pp.
69-113. Plenum Press, New York.

Carter, J. G. and G. R. Clark Il. 1985. Classification and phylogenetic
significance of molluscan shell microstructure. /n: Mollusks,
Notes for a Short Course, T. W. Broadhead, ed., organized by
D. J. Bottjer, C. S. Hickman and P. D. Ward. pp. 50-71. Univer-
sity of Tennessee, Department of Geological Sciences Studies
in Geology 13.

Crenshaw, M. A. and J. M. Neff. 1969. Decalcification at the mantle-
shell interface in molluscs. American Zoologist 9:881-885.

Fritz, L. W. and R. A. Lutz. 1986. Environmental perturbations reflected
in internal shell growth patterns of Corbicula fluminea
(Mollusca: Bivalvia). Veliger 28(4):401-417.

Hochachka, P. W. 1980. Living Without Oxygen; closed and open
systems in hypoxia tolerance. Harvard University Press, Cam-
bridge. 181 pp.

Lutz, R. A. and G. R. Clark. 1984. Seasonal and geographic varia-

tion in the shell microstructure of a salt-marsh bivalve [Geuken-
sia demissa (Dillwyn)]. Journal of Marine Research 42:943-956.

Lutz, R. A. and D. C. Rhoads. 1979. Shell structure of the Atlantic
ribbed mussel, Geukensia demissa (Dillwyn): A reevaluation.
Bulletin of the American Malacological Union for 1978:13-17.

Lutz, R. A. and D. C. Rhoads. 1980. Growth patterns within the
molluscan shell: in: Skeletal Growth of Aquatic Organisms, D.
C. Rhoads and R. A. Lutz, eds. pp. 203-254. Plenum Press,
New York.

Prezant, R. S. and A. Tan Tiu. 1986. Spiral crossed-lamellar shell
growth in the bivalve Corbicula. Transactions of the American
Microscopical Society 105(4):338-347.

Prezant, R. S., A. Tan Tiu and K. Chalermwat. 1988. Shell microstruc-
ture and color changes in stressed Corbicula c.f. fluminea
(Bivalvia: Heterodonta). Veliger 31(3/4):236-243.

Rhoads, D. C. and R. A. Lutz. 1980. Skeletal records of environmen-
tal change. /n: Skeletal Growth of Aquatic Organisms, D. C.
Rhoads and R. A. Lutz, eds. pp. 1-19. Plenum Press, New York.

Tan Tiu, A. 1987. Influence of environment on shell microstructure
of Corbicula fluminea and Polymesoda caroliniana (Bivalvia:
Heterodonta). Doctoral Dissertation, University of Southern
Mississippi, Hattiesburg. 148 pp.

Tan Tiu, A. 1988. Temporal and spatial variation of shell microstruc-
ture of Polymesoda caroliniana (Bivalvia: Heterodonta).
American Malacological Bulletin 6(2):199-206.

Tan Tiu, A. and R. S. Prezant. 1987. Shell microstructural responses
of Geukensia demissa granosissima (Mollusca: Bivalvia) to con-
tinual emergence. American Malacological Bulletin 5(2):173-176.

Tan Tiu, A. and R. S. Prezant. 1988. Shell tubules in Corbicula fluminea
(Bivalvia: Heterodonta): Functional morphology and microstruc-
ture. Nautilus: In press.

Wada, K. 1960. Crystal growth on the inner shell surface of Pinctada
martensii (Dunker) |. Journal of Electron Microscopy 9(1):21-23.

Date of manuscript acceptance: 23 August 1988

THE FUNCTIONAL MORPHOLOGY OF THE ORGANS OF THE
MANTLE CAVITY OF BATISSA VIOLACEA (LAMARCK, 1797)
(BIVALVIA: CORBICULACEA)

BRIAN MORTON
DEPARTMENT OF ZOOLOGY
THE UNIVERSITY OF HONG KONG
HONG KONG

ABSTRACT

Batissa violacea (Lamarck) occurs in rivers of the tropical Indo-West Pacific. It superficially
resembles members of the Unionidae but morphologically is clearly corbiculid. Like Polymesoda, Batissa
exhibits pedal gape feeding and can dig to considerable depths, probably to avoid desiccation. B. violacea
is dioecious, grows to 150 mm in shell length and is probably long-lived. The Corbiculidae are con-
sidered recent immigrants to fresh waters. A relatively unspecialized morphology and reproductive
strategy but with physiological and behavioural specializations to avoid drought have allowed this.
Reproductive specialisation characterises Corbicula, occupying river head-waters and lentic systems.
Batissa and Polymesoda thus provide living examples of how fresh waters have been colonised by

the Corbiculidae.

The Corbiculacea is a group of fresh or brackish water
heterodonts, mostly tropical in their distributions. Most interest
is with Corbicula, particularly the exclusively freshwater C.
fluminea (Muller) that has been spread artificially from its Asian
range to North and South America and Europe (Morton, 1986).
In North America it is a pest of power station cooling systems
which it clogs, although other problems have been en-
countered such as the blockage of drainage canals
(McMahon, 1983). Species of Polymesoda in both the western
Pacific and western Atlantic are similarly tropical and occur
in salt marshes and mangroves of estuaries (Morton, 1984).
They too have elicited interest for their physiological adapta-
tions to the harsh high intertidal (Depledge, 1985).

Prerequisite to an understanding of the species of Cor-
biculacea is anatomy, especially since the representatives of
this group appear to be phenotypically highly variable, e.g.
C. fluminea (Britton and Morton, 1986; Morton, 1987a). Few
authors have reported details of corbiculid anatomy. Prashad
(1920) described briefly the gross anatomy of Corbicula
fluminalis (Muller) and Dinamani (1957) that of Villorita
cyprinoides (Gray). Details of the anatomy of C. fluminea
are reported upon by Kraemer (1978, 1979, 1981), Kraemer
and Lott (1978) and Britton and Morton (1982). Morton (1976)
has described the anatomy of Polymesoda (Geloina) erosa
(Solander). There have been few studies of the genus Batissa,
most information resulting from geographic collections.
Raj and Fergusson (1980), however, have investigated the

osmolarity and ionic composition of the blood of Batissa
violacea (Lamarck, 1818) from Fiji, while Djajasasmita and
Budiman (1984) have presented some information on the
population density of this species in the Pisang River,
Sumatra.

The anatomy of this little known bivalve has not been
studied. This investigaton helps to remedy this situation but
also points out some interesting behavioural adaptations that,
linked with anatomical modifications, provides clues as to how
the colonisation of freshwaters by this important group of
bivalves has been achieved.

MATERIALS AND METHODS

Market specimens of Batissa violacea (60 - 90 mm shell
length) were first examined alive in Fiji and then subsequently
in Hong Kong. Following dissection, ciliary currents were
elucidated using fine carborundum and carmine. For
histological purposes, two specimens were removed from their
valves and fixed in aqueous Bouin’s fluid and, following
routine procedures, serially sectioned at 6 »m and alternate
slides stained in either Ehrlich’s haematoxylin and eosin or
Masson’s trichrome.

BIOLOGY

Batissa violacea is a tropical freshwater corbiculid
distributed throughout the western Pacific, i.e. Malaysia,

American Malacological Bulletin, No. 7(1) (1989):73-79

73

74 AMER. MALAC. BULL. 7(1) (1989)

Philippines, New Guinea, N.W. Australia and various Pacific
islands. Bentham-Jutting (1953) records it as the only species
from Java and from New Guinea (Bentham-Jutting, 1963);
Had (1976) records it from Fiji; McMichael (1967) from north-
western Australia. Brandt (1974) records B. similis Prime, 1860
from the Nicobar Islands and rivers in Thailand.

Raj and Fergusson (1980) reported upon the osmolarity
(64.1 + 8 m Osmol) and ionic composition of the blood of
Batissa violacea (Na* , 35; K+, 898; Ca2*+, 2.98; Mg2*,
26.08; Cl-, 4.26 mmol kg"! water) and showed them to be com-
parable to the better-known unionids Anodonta cygnea Lin-
naeus (42 m Osmol; Nat , 5.3; Kt, 21.3; Ca2+ , 12.0;Mg2*,
4.5; Cl-, 2.4 mmol kg"! water) and Hyridella menziesi Gray
(62+ 15 m Osmol; Nat, 1.6; K+, 7.2; Ca2+, 2.0; Cl, 13.7
mmol kg"! water). They concluded that B. violacea is the result
of a relatively recent immigration into fresh waters by the Cor-
biculidae, a view upheld by this author (Morton, 1985; 1987b).
Djajasasmita and Budiman (1984) demonstrates that the
species occurs in the muds of the banks and river beds in
Indonesia.

Eight specimens of Batissa violacea obtained alive
from Fiji were placed into a freshwater aquarium at 21°C in
Hong Kong, with 15 cm depth of sand. They burrowed rapid-
ly to the full depth. This was thought to be a possible response
to perceived drying. Subsequently, two individuals took up
residence at the sand water interface, only the tips of their
shells above it with the siphons projecting from between the
valve margins only slightly. Six other specimens, however, re-
mained buried to a depth of ~ 10cm from the posterior edge
of the shell and even with periodic removal, returned to this
depth.

FUNCTIONAL MORPHOLOGY

SHELL

The shell of Batissa violacea is equivalve and approx-
imately equilateral, although the posterior is somewhat
elongated in relation to the anterior (Fig. 1A). The outline
varies considerably, some shells being more rounded, others
elongated. A maximum length of 150 mm has been record-
ed (Bentham-Jutting, 1953). The periostracum is thick and

A

dark; younger shells appear dark vioiet, older ones black. The
older parts of the shell, around the umbones, are often erod-
ed. The opisthodetic external ligament is large. The posterior
margin is sometimes square and an umbonal ridge extends
to its postero-ventral border much as in Polymesoda erosa
(Morton, 1976). Seen from the anterior, the shell is narrow,
the widest region, as in most burrowers, being dorsal to the
mid point of the dorso-ventral axis of the shell (Fig. 1B: x-x).
The shell margin gapes anteriorly (Fig. 1B).

The nacreous internal surface of the shell is varying
shades of purple, particularly external to the pallial line. This
is recessed deeply from within the shell margin and is dou-
ble, being composed of an inner and an outer line [Fig. 2:
PL(I), PL(O)]. The latter is much thicker than the former.
Posteriorly, there is a shallow pallial sinus (PS). The anterior
and posterior adductor muscles (AA, PA) are deeply
impressed into the shell of older individuals, just below the
large hinge plate.

The massive external ligament comprises a posterior
outer ligament layer (POL) and an inner ligament layer (IL)
(Yonge, 1978); if there is an anterior outer ligament layer it
is either lost or very reduced. The ligament is overlain by
periostracum (PE), extending beyond the ligament as a thick
wad that covers the shell and darkens the inner margin of
each valve.

The hinge plate (Fig. 3) is broad and the left valve
possesses three cardinal teeth (CT), posteriorly directed.
These interlock with two teeth in the right valve. The central
cardinal tooth of the right valve can be bifid. The left valve
has anterior and slightly longer posterior lateral teeth (LT) that
fit into sockets on the right valve (LTS). The lateral teeth of
the left valve and the lower lips of the sockets of the right valve
are grooved transversely.

ADDUCTOR MUSCLES

Batissa violacea is approximately isomyarian, the two
adductors being located under the ends of the hinge plate
(Fig. 2: AA, PA). Also under each hinge plate beneath the
lateral teeth and internal to each adductor is a pedal retrac-
tor muscle (APR, PPR). The adductor muscles are divided

BOO

Sd

5cm

Fig. 1. Batissa violacea. The shell as seen from A, the right side and B, from the anterior, showing (x--x) the maximum width.

MORTON: MANTLE CAVITY ORGANS OF BATISSA 75

IL POL

PL(I)

Fig. 2. Batissa violacea. An internal view of the left shell valve. [AA,
anterior adductor muscle scar; APR, anterior pedal retractor muscle
scar; IL, inner ligament layer; PA, posterior adductor muscle scar;
PE, periostracum; PL(I), inner component of the pallial line; PL(O),
outer component of the pallial line; POL, posterior outer ligament
layer; PPR, posterior pedal retractor muscle scar; PS, pallial sinus;
U, umbo].

into slow and quick components of smooth and striated mus-
cle bundles.

SIPHONS

The siphons (Fig. 4) are located posteriorly, the ex-
halant (ES) being small and fringed apically by small papiilae.
The inhalant (IS) is larger and crowned by a complex array
of papillae. There are usually 24 large primary papillae, in-
terspersed by an equal number of smaller secondary papillae.
These are interspersed by some 48 tertiary papillae and these,
in turn, by approximately 96 tiny quaternary papillae. The
siphonal apparatus is thus well endowed with sensory papillae
and the shallowness of the pallial sinus attests to the fact that
the siphons can extend only slightly from between the shell
valves. The siphons are dark brown and flecked with yellow.
The inner reaches of the exhalant siphon are yellow. The
siphons are formed by fusion of the inner mantle folds only
and are thus of type A (Yonge, 1957, 1982). Papillae around
the base of the siphons extend dorsally and ventrally as
gradually merging and diminishing rows, the latter towards
the pedal gape (PG).

MANTLE MARGIN

The mantle margin comprises the usual three folds
(Fig. 5), except that the inner fold is divided into two com-
ponents: inner [IMF(I)] and outer [IMF(O)]. The inner com-
ponent contains the groove of a major rejectory tract (RT), the
outer component has sensory papillae. The inner fold con-
tains the inner component of the pallial retractor muscle
[PRM(I)], arising from the inner pallial line [Fig. 2: PL(I)] and
constituting the major muscles of the mantle margin where
left and right lobes fuse, i.e. between inhalant and exhalant

siphons and inhalant siphon and pedal gape. The inner fold
contains a few, basiphilic sub-epithelial, mucous cells. The
middle fold (MMF) is relatively small and forms the surface
against which the periostracum (P) is secreted from the in-
ner surface of the outer fold. The larger outer component of
the pallial retractor muscle [PRM(O)], arising from the outer
component of the pallial line [Fig. 2: PL(O)], is closely
associated with the periostracal groove. The outer mantle fold
(OMF) is large and contains a haemocoel. There is a pallial
nerve (PN) between inner and outer components of the pallial
retractor muscle.

PEDAL GAPE

The anterior pedal gape (Fig. 6: PG) is long, the inner
folds forming it being extensively equipped with blunt-tipped
papillae. These are larger at the posterior and anterior ex-
tremities of the pedal gape. Anteriorly too, the shell is
emarginated (Fig. 1B) and water is forcibly expelled here when
the animal is handled.

CTENIDIA

The ctenidia are flat, homorhabdic, eulamellibranch
and plicate. Each plica comprises up to 48 filaments. Each
ctenidium comprises inner and outer demibranchs (Fig. 6A:
ID, OD), the latter dorsoventrally much shorter than the former.

The ctenidia are relatively small occupying, approx-
imately, the postero-dorsal quadrant of the mantle cavity. The
ctenidial ciliation is of type C(2) (Atkins, 1937a) (Fig. 6B), also
seen in Polymesoda erosa (Morton, 1976). Acceptance tracts
are thus located in the ventral marginal food grooves of both
demibranchs and in the ctenidial axis, but not in the junctions
created by the ascending lamellae of the inner and outer demi-
branchs with the visceral mass and mantle respectively. The

CT
CT nae

LT

Fig. 3. Batissa violacea. The hinge plate, left valve above, right below
(CT, cardinal tooth; LT, lateral tooth; LTS, lateral tooth socket).

76 AMER. MALAC. BULL. 7(1) (1989)

edges of the ascending lamellae of both inner and outer
demibranchs are connected to the visceral mass and mantle
respectively by tissue fusions (Atkins, 1937b).

LABIAL PALPS

The labial palps (Fig. 6: LP) are large and their
posterior edges are associated closely with the ventral
marginal food grooves of the ctenidia. The ctenidial-labial palp
junction is of Category 3 (Stasek, 1963). Thus, material arriv-
ing at the ctenidial termini move on to a short distal oral
groove, but is probably collected from the ventral food grooves
by the general palp surfaces before reaching this point.

Fig. 4. Batissa violacea: The posterior shell margin showing the
siphons (ES, exhalant siphon; IS, inhalant siphon; PG, pedal gape).

PRMI(I)
RT

PRMIO)

PN

IMF(O)

MMF

OMF

a |
250 um

Fig. 5. Batissa violacea. A transverse section through the right man-
tle lobe at the pedal gape [IMF(I), inner component of the inner mantle
fold; IMF(O), outer component of the inner mantle fold; MC, mucous
cell; MMF, middle mantle fold; OMF, outer mantle fold; P,
periostracum; PN, pallial nerve; PRM(I), inner component of the pallial
retractor muscle; PRM(QO), outer component of the pallial retractor
muscle; RT, rejectory tract].

The detailed ciliary currents of two palp ridges and an
intervening groove are shown in figure 7. Fine, accepted par-
ticles are transported rapidly over the surface of the palps
to the distal oral groove and thence via a short proximal oral
groove to the mouth, located ventral to the anterior pedal
retractor muscle. Large, unwanted particles fall into the depths
of the groove and are transported towards the ventral edge
of the palp where they then pass to its free tip and are re-
jected. On the oral and aboral faces of the palp ridges are
complex re-sorting currents that allow Batissa violacea to ac-
cept or reject intermediate-sized particles in greater or lesser
quantities.

CILIARY CURRENTS OF THE VISCERAL
MASS AND MANTLE

The ciliary currents of the visceral mass are shown in
figure 6. At the approximate junction of the foot (F) with the

MORTON: MANTLE CAVITY ORGANS OF BATISSA TEE

OM
it mi iD Ne

LP

VM

Fig. 6. Batissa violacea. A, The animal as seen from the right side and after removal of the right shell valve and mantle lobe. Ciliary currents
are indicated by arrows; B, a diagrammatic transverse section through a single ctenidium showing the ciliary currents and acceptance tracts
(¢). (AA, anterior adductor muscle; ES, exhalant siphon; F, foot; H, heart; ID, inner demibranch; IS, inhalant siphon; K, kidney; LP, labial palp;
OD, outer demibranch; PA, posterior adductor muscle; PG, pedal gape; PPR, posterior pedal retractor muscle; VM, visceral mass).

visceral mass (VM), a rejectory tract extends along each side
from the position of the palp tip to the posterior edge of the
visceral mass. These tracts are fed from above by downward
cleansing currents and from below by cilia collecting material
from the dorsal regions of the foot. The foot itself is free of
such cleansing cilia. Material collected in the left and right
grooves eventually falls onto the mantle below.

The ciliary currents of the mantle complement those
of the visceral mass. Each lobe possesses a deep rejection
tract formed at the junction of the inner component of the in-
ner mantle fold with the general mantle surface. Unwanted
material from the general mantle surface is passed
downwards into the left and right rejectory tracts and
transported towards the base of the inhalant siphon where
it is rejected as pseudofaeces (Fig. 6).

ORGANS OF THE PERICARDIUM

The heart (Fig. 8) lies beneath the ligament. It com-
prises a single ventricle (V) penetrated by the rectum (R), and
a pair of auricles (AU). From the posteroventral edge of the
pericardium arise a pair of renopericardial apertures (RPA)
that open into the distal limbs of the kidneys (K). The kidneys
are located between the pericardium and the posterior ad-
ductor muscle (PA). The rectum passes above them. The
proximal limbs of the kidneys open into the suprabranchial
chamber, as renal apertures (RA), between the ctenidial axis
and the point of attachment of the ascending lamella of the
inner demibranch to the visceral mass (PALID). Located close

Fig. 7. Batissa violacea. A diagrammatic representation of two ridges
of a labial palp to show the various ciliary tracts (for explanation see
text).

78 AMER. MALAC. BULL. 7(1) (1989)

Fig. 8. Batissa violacea. The organs of the pericardium as seen from
the right side. (A, anus; AU, auricle; CA, ctenidial axis; G, gonopore;
K, kidney; PA, posterior adductor muscle; PALID, point of attachment
of ascending lamella of the inner demibranch to the visceral mass;
PEG, pericardial gland; PPR, posterior pedal retractor muscle; R,
rectum; RA, renal aperture; RPA, renopericardial aperture; V,
ventricle).

to the renal apertures are the gonopores (G). Batissa violacea
is dioecious. Eggs shed in the laboratory measured between
80-120 um. The pericardial gland (PEG) is largely associated
with the anterior pericardium (White, 1942) as in Polymesoda
(Geloina) erosa (Morton, 1976).

DISCUSSION

On first inspection, the black shell of Batissa violacea
is strongly reminiscent of riverine Unionidae (see illustrations
in Burch, 1975) and indeed this species, among the Cor-
biculidae, is a tropical riverine species (Djajasasmita and
Budiman, 1984). The similarity in shell form can be regarded
as an example of convergent evolution.

In other aspects of its anatomy, however, Batissa
violacea is a typical corbiculid, similar in most respects to
Polymesoda (Geloina) erosa (Morton, 1976). An important
feature of the latter species is that the pallial line is single,
whereas in B. violacea it is double. The condition in Batissa
stems from duplication of the inner mantle fold and a close
association between the outer pallial retractor component with
the periostracal groove. The periostracum of B. violacea is
thick, and could have an important protective function for the
shell in acidic, tropical waters.

A further important feature of the Batissa violacea shell
is the anterior gape through which water is ejected when the
animal is handled. This is also seen in Polymesoda erosa,
which has been shown to feed from subterranean water (Mor-
ton, 1976). This probably also occurs in B. violacea, and it
is perhaps significant that this species is capable of living
deep within the sediment with no siphonal access to the
substratum surface. The posterior margin of the shell would
generally be located at the sediment-water interface but in
times of drought the species can probably dig deeper into
the sediment and effect simple exchange with the water table

via the pedal gape. It is interesting to note that Sinclair and
Isom (1963) illustrate Corbicula fluminea as capable of living
in deep, moist, sediments. Indeed, the species has been dug
up alive from sands with little evidence of flowing water, and
has been found to cause problems subsequently in concrete
aggregates.

McMahon (1983) has reviewed mechanisms of desic-
cation tolerance in Corbicula fluminea and shown that aerial
respiration is possible for a period of a few days (McMahon,
1979) at the posterior mantle margin, as in Polymesoda erosa
and P proxima (= P expansa) (Morton, 1975, 1976, 1984). With
the corbiculid capacity for pedal gape feeding and the poten-
tial for deep residence, it would seem that the first response
to surface drying is to dig to deeper moisture levels. Highly
stressed C. fluminea, however, crawl to the surface and are
washed downstream (McMahon, 1983). Prezant and Chalerm-
wat (1984) report that C. fluminea produces mucous drogue
lines that facilitate downstream floatation. The first facility could
be important in the relatively recent colonisation of freshwaters
by the Corbiculidae, the latter two particular behaviours ex-
pressed under conditions of stress by C. fluminea, a head-water
and lentic species. Derived from a marine ancestor (Raj and
Fergusson, 1980; Morton, 1987b), modern representatives of
the Corbiculidae have had to withstand periodic drought,
either tidal in Polymesoda (Morton, 1976) or seasonal as in
Batissa and Corbicula. The mechanism to overcome drought
now demonstrated for Batissa as well as Polymesoda points
to an important behavioural adaptation that has facilitated col-
onization of first estuarine and then lacustrine habitats.

In all other respects, the Corbiculidae are little different
morphologically from other Veneroida. Like Polymesoda (Mor-
ton, 1985), Batissa is dioecious and non-incubatory and, with
a maximum shell length of 150 mm and numerous growth
lines (Bentham-Jutting, 1953), probably long-lived. The
osmolarity and ionic composition of Batissa blood indicates
recent colonisation of freshwaters (Raj and Fergusson, 1980).
A simple morphology and reproductive strategy are also sug-
gestive of this. Thus, physiological specializations and
behavioural adaptations to avoid drought were critical for the
exploitation of freshwaters particularly the lower reaches of
rivers. Later reproductive specialisations, i.e. a variable sex
ratio and incidence of hermaphroditism, as in Corbicula
flumina, allowed colonisation of river head waters and lentic
systems, with reductions in size and longevity (Morton, 1987b).
Behavioural specialisations to avoid drought (McMahon, 1983)
were, however, important in the progressive colonisation of
fresh waters by the Corbiculidae.

ACKNOWLEDGMENTS

Initial research on Batissa violacea was undertaken at the
University of the South Pacific, Fiji, in January 1985 whilst the author
was holder of an Association of Commonwealth Universities Senior
Travelling Fellowship sponsored by the Leverhulme Trust. | am grateful
to the Trust and to Dr. Uday Raj (University of the South Pacific) for
much help and hospitality. | subsequently received further live
specimens of B. violacea through the courtesy of Ms. Milika Naqasima
(University of the South Pacific). | am also grateful to Mr. H. C. Leung
(University of Hong Kong Kong) for histological assistance.

MORTON: MANTLE CAVITY ORGANS OF BATISSA 79

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Date of manuscript acceptance: 22 November 1988

Fa
f

BIVALVES IN THE GENUS CORBICULA (BIVALVIA: CORBICULIDAE)
IN THE SOVIET UNION WITH A CATALOGUE OF TYPE MATERIALS
IN THE ZOOLOGICAL INSTITUTE, ACADEMY OF SCIENCES
OF THE U.S.S.R., LENINGRAD

CLEMENT L. COUNTS, Ill
COASTAL ECOLOGY RESEARCH LABORATORY
UNIVERSITY OF MARYLAND EASTERN SHORE
PRINCESS ANNE, MARYLAND 21853, U.S.A.

ABSTRACT

Type materials for three species and three subspecies of bivalves in the genus Corbicula held
in the collections of the Zoological Institute, Academy of Sciences of the U.S.S.R., Leningrad, are reported.
Catalogue numbers, number and type of specimens, locality data from collection labels, and dates
of collection are provided for 39 lots of type materials for C. ferghanensis, C. fluminea extrema, C. fluminea
praebaicalensis, C. lindholmi, C. suifuensis and C. suifuensis finitima. Notes on other species of fossil
and Recent Corbicula described from the U.S.S.R. are given with a discussion of their zoogeography.
The debate on the number of species of Corbicula within the Soviet Union is discussed with a review
of current practices used to resolve this systematic problem.

Bivalves in the genus Corbicula Muhlfeldt, 1811, have
been the object of malacological study in the Soviet Union
for many years. While some research on the morphology,
physiology and ecology of corbiculids has been conducted
by Soviet malacologists (Mitropolskie, 1963; Butenko, 1967;
Sultanov et al., 1972; Kasymov and Gadshiyeva, 1974; Alimov,
1974, 1975; Karpevich, 1975; Yaroslavteva et al., 1981; Zaiko
and Romanenko, 1981; Komendantov, 1984), most reports on
Soviet corbiculids concern their paleontology (Androussov,
1923; Slokedewitsch, 1938; Otatume, 1943; Suzuki, 1943;
Volkova, 1962; Andrusov, 1966; Krylova, 1966; Yakushima,
1968, 1973; Zhubkova et al., 1968; Ibadov, 1972; Dubinovs’kyi
et al., 1974; Khubka, 1979). This is not so surprising since most
Soviet malacologists are trained as geologists with an em-
phasis on stratigraphy and paleontology (Amitrov, 1983;
Counts, 1986). There have also been some reports on the
biogeography of bivalves in the genus Corbicula within the
Soviet Union (Rosen, 1914; Sidaroff, 1929; Decksbach, 1943;
Zhadin, 1952; Aliev, 1960; Volkova, 1962; Mitropolskie, 1963;
Kasymov, 1972; Kasymov and Gadshiyeva, 1974; Karpevich,
1975; Izzatullaev, 1980; Izzatullaev and Starobogatov, 1985).
Two recent reviews of the systematics of genus Corbicula in
the Soviet Union have appeared (Kursalova and Starobogatov,
1971; Izzatullaev, 1980), one of which (Kursalova and
Starobogatov, 1971) included descriptions of new taxa.

Several species and subspecies of recent and fossil
bivalves in the genus Corbicula have been described from

Soviet waters over the past century (von Martens, 1874;
Clessin, 1879; Androussov, 1923; Lindholm, 1927; Slokede-
witsch, 1938; Otatume, 1943; Suzuki, 1943; Popova, 1968;
Yakushima, 1968; Zhubkova et al., 1968; Kursalova and
Starobogatov, 1971). The type materials of three species and
three subspecies are located in the collections of the
Zoological Institute, Academy of Sciences of the U.S.S.R., Len-
ingrad. This paper discusses the species of bivalves in the
genus Corbicula in the Soviet Union and presents notes on
the type materials held in the Zoological Institute’s collections.

TYPE MATERIALS

A survey of the corbiculid materials in the collections
of the Zoological Institute of the Academy of Sciences of the
U.S.S.R., Leningrad, was made during April 3 - 16, 1986. Due
to the institutional practice of providing only lots selected from
the card catalogue, it was not possible to examine the entire
collection in the ranges.

The type material for three species and three
subspecies of bivalves in the genus Corbicula now in the col-
lections of the Zoological Institute of the Academy of Sciences
of the U.S.S.R., Leningrad (AH-CCCP) are presented below.
All of these species were described from Soviet waters. It
should be noted that the rules for designation of type
specimens and localities are somewhat more flexible among
Soviet malacologists than among their western colleagues.

American Malacological Bulletin, Vol. 7(1) (1989):81-86

81

82 AMER. MALAC. BULL. 7(1) (1989)

In some instances, specimens designated as paratypes are
from localities other than the holotype or syntypes. Further,
some paratype series were collected over a period of 40 years.
While these idiosyncracies are not unique to materials in the
collections of the AH-CCCP, they do apply to all the type
materials referred to the genus Corbicula in that institution.

None of the type materials discussed below were
designated by lot number in the literature that describes them
with the exception of the holotype of Corbicula fluminalis
praebaicalensis (Popova, 1968). Taxa, catalogue numbers,
number of specimens, type localities, and dates of collection
are provided below.

Corbicula ferghanensis Kursalova and Starobogatov, 1971,
p. 95 (Uzbek S.S.R., Aral Sea Region, Ferghana River).

AH-CCCP 446-1961, No. 1, Holotype, Ferghana (River), 4 IV
1931. Collected dead.

AH-CCCP 446-1961, No. 2, 10 + 1/2 paratypes, dry, Ferghana
(River), 4 IV 1931. Collected dead.

AH-CCCP 446-1961, No. 3, 2 + 6/2 paratypes, dry, Ferghana
(River), 1931.

AH-CCCP 446-1961, No. 4, 8 paratypes, dry, Ferghana (River),
near (power?) station, 13 IV 1931.

AH-CCCP 446-1961, No. 5, 2/2 paratypes, dry, Karakalpak
Autonomous S.S.R., Muynak, 26 VI 1947.

AH-CCCP 41-1964, No. 6, 1 paratype, dry, Farkhskoe Reser-
voir and tributaries, Tadzhikistan (Tadzhik S.S.R.), 14
VII 1960.

AH-CCCP 41-1964, No. 7, 1 paratype, dry, shallow water,
Kairak-Kumskoe Reservoir, Tadzhikistan (Tadzhik
S.S.R.), 5 X 1960.

AH-CCCP 41-1964, No. 8, 1 paratype, dry, Kairak-Kumskoe
Reservoir, Tadzhikistan (Tadzhik S.S.R.), 30 XI 1960.

AH-CCCP 483-1967, No. 9, 37 + 29/2 paratypes, dry, Right
bank of Syr-Dar’ya River, Samgar Canal from Kairak-
Kumskoe Reservoir, 25 VII 1967.

AH-CCCP 284, No. 10, 3/2 paratypes, dry, (no locality, no date).

AH-CCCP 284-1969, No. 11, 9/2 paratypes, dry, (no locality),
1968.

AH-CCCP 175-1929, No. 12, 1/2 paratype, dry, (no locality),
21-22 IX 1928.

AH-CCCP 241-1962, No. 13, 1/2 paratype, dry, (no locality, no
date).

AH-CCCP 359-1935, No. 14, 2/2 paratypes, dry, (no locality,
no date).

AH-CCCP 452-1973, No. 15, 4/2 paratypes. Quaternary fossils,
(no locality, no dates). The label accompanying these
specimens indicates they are also paratypes of Cor-
bicula fluminea praebaicalensis.

Remarks: Corbicula ferghanensis is also reported from

a large irrigation ditch off the Ferghan River near the village

of Arkangelisk (Kursalova and Starobogatov, 1971). Izzatullaev

(1980) reported C. ferghanensis in the Syr Dar’ya basin

(Ferghan River) and Amu Dar’ya basin (Karakum and

Samarkand) in the Tadzhik and Uzbek S.S.R. He also noted

its presence in the environs of lakes Baikal, Irtash, and

Balkash.

Corbicula fluminea extrema Lindholm, 1927, 28:550.

AH-CCCP 465-1929, No. 2, 4 syntypes in alcohol, Amur
estuary near Dzhaore, 19 VI 1928.

AH-CCCP 198-1961, No. 2, 2 syntypes, dry, Amur estuary near
Dzhaore, 19 VI 1928.

AH-CCCP 465-1929, No. 3, 2 + 2/2 syntypes, in alcohol,
Osmrov Canal, 18 VIII 1928.

AH-CCCP 465-1929, No. 4, 5 syntypes, dry, Vladimir Bay, Sea
of Japan, VIII 1927. (All collected dead with umbones
and internal shell features badly eroded).

AH-CCCP 465-1929, No. 5, 1 syntype, dry, Sachalin (Sakhalin)
Island near Astrakhanovskii, 12 VII 1928.

AH-CCCP 456-1929, No. 6, 13 syntypes, dry, Amur estuary
near Dzhaore, 26 VI 1928.

Remarks: Kursalova and Starobogatov (1971) recog-
nized Corbicula fluminea extrema (Corbicula fluminalis extrema
in their paper) as a junior synonym of C. japonica Prime, 1864.
They reported the species to be present in the waters of the
continental coast of the Sea of Japan, the Amur River estuary,
and the southern Kurile Islands and Sakhalin Island, as well
as Japan.

Corbicula fluminea praebaicalenis Popova, 1968, pp. 257-258,
Pl. 1, Figs. 13-15 (northwest Prebaikal, River Anga).

AH-CCCP 452-1973, No. 1, 22 /2 paratypes, Quaternary fossils,
(no locality, no date).

AH-CCCP 452-1973, No. 2, 4/2 paratypes, Quaternary fossils,
(no locality, no date). Reidentified as Corbicula
ferghanensis (AH-CCCP 452-1973, No. 16).
Remarks: The holotype of Corbicula fluminea prae-

baicalensis is located in the collection of the Limnological In-

stitute, Academy of Sciences of the U.S.S.R., Irkutsk (No.
20/64A) (designated by Popova, 1968). Kursalova and

Starobogatov (1971) report the taxon to be a junior synonym

of C. tibetensis (Prashad, 1929).

Corbicula lindholmi Kursalova and Starobogatov, 1971, p. 94.

AH-CCCP 205, 1938, No. 1, Holotype, Ussuri River, Suifun
River (23-25 Vil 1924).

AH-CCCP 460-1929, No. 2, 1 paratype in alcohol, Suifun River
and its estuary, 20 VII 1925.

AH-CCCP 198-1961, No. 3, 2 paratypes, dry, Suifun River, 18
VIIl 1928. (Three specimens are listed on the label but
only two specimens are in the lot. Both were collected
dead and show erosion of the internal shell features,
especially the lateral teeth.)

AH-CCCP 198-1961, No. 4, 1 paratype, dry, Suifin River, 1925.

AH-CCCP 212-1925, No. 5, 1 paratype, dry, (no locality), 1925.
Remarks: Kursalova and Starobogatov (1971) also

report Corbicula lindholmi from the South Primorie and up-

per portion of the Sungari River basin, the lower portion of
the Pauecheza River near the villages of Dulakeet and

Derzharena. They noted specimens were cast ashore on Slav-

yank Creek (also known as Velik Creek) at Razina. They also

report populations near the frontier of the People’s Republic
of China at the village of Velikovsk.

COUNTS: CORBICULA IN THE SOVIET UNION 83

Corbicula suifuensis Lindholm, 1925, p. 29; 1927, Pl. 32,
Figs. 1a, 1b (Suifun River near Razhdolnaya, south-
eastern Siberia).

AH-CCCP 216-1924, Holotype, dry, Suifun River, VII 1924.

AH-CCCP 216-1924, No. 2, 1 paratype, dry, Suifun River, VII
1924.

AH-CCCP 205-1938, No. 3, 3 paratypes, dry, Suifun River,
23-25 VII 1942.

AH-CCCP 205-1938, No. 4, 1 paratype, dry, Suifun River, VII
1924 (‘‘C. producta’’ is written on the valves).
AH-CCCP 460-1929, No. 6, 1 paratype, dry, Suifun River and

its estuary, Primorie region, (no date).

AH-CCCP 460-1929, No. 7, 1 paratype, dry, Suifun River and
its estuary, Primorie region, (no date).

AH-CCCP 460-1929, No. 8, 1 paratype, dry, Suifun River and
its estuary, Primorie region, (no date).

AH-CCCP 460-1929, No. 9, 1 paratype, dry, Suifun River and
its estuary, Primorie region, (no date).

AH-CCCP 460-1929, No. 10, 7 paratypes, dry, Suifun River
and its estuary, Primorie region, (no date).
Remarks: Other records are the River Maikhe, coast

of Posaeta Bay, Primorie (Kursalova and Starobogatov, 1971).

Corbicula suifuensis finitima Lindholm, 1927, pp. 553-554,
Pl. 32, Figs. 2a, 2b (Estuary of the Mai-che River, southeastern
Siberia).

AH-CCCP 460-1929, No. 1, Holotype, dry, rivers and estuaries
of the Primorie region, (no date).

AH-CCCP 460-1929, No. 2, 2 paratypes, dry, rivers and
estuaries of the Primorie region, 24 VI 1924.
Remarks: Kursalova and Starobogatov (1971) report

Corbicula suifuensis finitima to be a junior synonym of C. elatior

Martens, 1905.

DISCUSSION

Other bivalve taxa described from waters of the Soviet
Union have been referred to the genus Corbicula. Clessin
(1879) described C. hohenackeri from the Jalysch River of the
Caucasus. The type materials for this species were in the
Stuttgart Museum and are believed to have been destroyed
during World War Il. Martens (1874) described C. minima from
Samarkand and Lindholm (1933) later reported collecting the
species in Central Asia. The type materials for the species
was believed to be in the collections of the Zoological Museum
in Moscow. However, all attempts by Soviet malacologists to
locate these materials have failed and the types are now con-
sidered lost (Starobogatov, pers. comm., 1986).

Several fossil species have been described from strata
in the Soviet Union. Androussov (1923) described Corbicula
fluminalis apscheronica from the Pleistocene of the Apscheron
Peninsula, Azerbaidzhan. C. fonsata and C. kovatschensis
were described from the Soviet Union Far East (Slokede-
witsch, 1938). Many fossil species have been described from
the strata of Sakhalin Island. These include C. sakakibarai
Otatume, 1943 from the Naibuchi Group, south Sakhalin

Island; C. gabliana lautenschlageri Zhubkova et al., 1968
described from the Pliocene of the River Tym; C. matachien-
sis Zhubkova et al., 1968 from the upper Miocene-Pliocene
of the River Mach; and C. glabiana adamensis described from
the upper Miocene-Pliocene of Sakhalin Island. Further south,
Yakushima (1968) described C. susaensis from the upper
Cretaceous of the south Primorye. My efforts to locate type
material for these taxa have not been successful. However,
type materials for two fossil taxa are presently in the collec-
tion of the Institute of Geology and Paleontology (IGPS) of
the University of Tokyo: C. shimizui Suzuki, 1943 (Holotype
IGPS 8353b, Paratype IGPS 8353a) and C. sachalinensis
Suzuki, 1943 (Holotype and Paratypes IGPS 8353a); both
species from the Tertiary Aquitanian Mach Group of the mid-
dle course of the Tumis River, North Sakhalin Island.

Corbicula fluminalis (Muller, 1774) appears to be the
most widely distributed species within the Soviet Union. This
is reflected both in published accounts and in the collections
of the AH-CCCP in so far as | was able to examine them. C.
fluminalis has been generally reported from the Caucasus in
Azerbaidzhan, Iran, Syria, Afghanistan, India, Soviet Central
Asia, and Baluchistan (Kasymov, 1972). Likharev and
Starobogatov (1967) and Solem (1979) have also reported C.
fluminalis from Afghanistan. Lindholm (1930) reported C.
fluminalis from Buchara (now Uzbek S.S.R.). Decksbach (1943)
reported C. fluminalis from Azerbaidzhan and Kazakhistan.
He also reported the species in Uzbekistan at the mouth of
the Amu Dar’ya River and in the Amur Basin. He further noted
its presence in Turkmenia in the valley of the Murgrab River.
Zhadin (1952) reported C. fluminalis to be distributed
throughout the bays of the southern Caspian Sea, the
Transcaucasus (Kura River basin and Lake Adzhikabul), in
the irrigation canals of Ashkabad and the lower reaches and
delta of the Amu Dar’ya at Samarkand, as well as the Murgab
River. He further noted that C. fluminalis are found as fossils
in Quaternary strata in the Moldavian S.S.R. (Dniester Ter-
races), the Ukranian S.S.R., and in the Pleistocene Apscheron
Layer of the Betekei River, western Siberia. Volkova (1962)
reported C. fluminalis from the lower reaches of the Irtysh
River. Kursalova and Starobogatov (1971) reported the species
in the Azerbaidzhan and Turkmen S.S.R. and in the Amu
Dar’ia. This may be the species referred to by Sidaroff (1929)
in the Aral Sea. Boettger (1881) reported the subspecies C.
fluminalis crassula ‘Mousson’ Bellardi, 1854 and C. fluminalis
compressa ‘Mousson’ Deshayes, 1854 from Lake Adzhikabul,
Transcaucasus (Azerbaidzhan S.S.R.).

Corbicula japonica Prime, 1864, is also reported to be
widely distributed in the Soviet Far East. This, again, is
reflected in both published accounts and in the records of the
AH-CCCP. C. japonica is variously reported from the
Razhdolnaya River (Zaiko and Romanenko, 1981) and from
the brackish water reaches of the Amur River estuary and
from Dzhaore Cape, Uarke Cape, the Chastye Islands of
Sakhalin Island, and in the northwestern part of the estuary
at Schatije Bay and Khalesovo Cape (Garkalina and
Moskvicheva, 1984). Kursalova and Starobogatov (1971) note
that C. japonica is widespread throughout the continental
estuaries of the Sea of Japan, the Amur estuary, Sakhalin

84 AMER. MALAC. BULL. 7(1) (1989)

Island, and the Kurile Islands. Other records for C. japonica
in Soviet waters from AH-CCCP include: Sakhalin Island on
a coastal spit near Nabil; Lake Ain, Sakhalin Island; Sugan
River Bay; Amurskiy Bay near Ussi; the Sea of Okhotsk,
Sakhalin Zaliv, and the northern limits of Amurskiy Bay.

Soviet malacologists are debating the number of
species referable to the genus Corbicula within the Soviet
Union (Kasymov, 1972; Izzatullaev, 1980; Izzatullaev and
Starobogatov, 1985). These arguments take on many of the
features of the same debate occurring in North America con-
cerning the number of species of corbiculids in that continent
(Britton and Morton, 1979, 1986; Hillis and Patton, 1980;
McLeod and Sailstad, 1981; McLeod, 1986; Morton, 1987). In
the United States, this debate has been resolved to the
satisfaction of most malacologists by the use of biochemical
genetic techniques (although there now appears to be a
debate about whether there is a debate). Soviet malacologists
are attempting to resolve their problems using morphological,
ecological, and reproductive characteristics (Izzatullaev, 1980).

Kursalova and Starobogatov (1971) reported fourteen
species of Corbicula within north and west Asia and Europe.
These were C. japonica, C. elatior, C. producta Martens, 1905,
C. finitima, C. lindholmi, C. fluminalis, C. cor (Lamarck, 1818),
C. consobrina (Caillaud, 1826), C. delessertiana Prime, 1870,
C. pusilla (Philippi, 1846), C. purpurea Prime, 1864, C. hebraica
Locard, 1883, C. tibetensis, and C. ferghanensis. These
species were identified on the basis of shell characters with
particular emphasis on tooth morphology.

Izzatullaev (1980) identified five species of corbiculid
bivalves from the Central Asian republics on the basis of their
reproductive biology. These included Corbicula cor, C.
fluminalis, and C. purpurea which were reported to be
oviparous. C. tibetensis and C. ferghanensis were referred to
the Australian genus Corbiculina Dall, 1903 on the basis of
their ovoviviparity.

Soviet malacologists regard the curvature (or degree
of inflation) of the shell (Logvinenko and Starobogatov, 1971)
as an important systematic tool in identifying their corbiculid
species (e.g. Izzatullaev, 1980). The method involves tracing
the curvature of a valve using a camera lucida and matching
the resulting curve to other specimens. It should be noted that
this method does not stand as a single test for taxon assign-
ment and that other shell characters are used. However, my
experience indicated that the shell curvature method was used
in many cases as the critical determining factor in referring
corbiculids to taxa in the Soviet Union. The most troublesome
aspect of depending upon the shell curvature is that it does
not take into account the physical and biological factors that
can affect interpopulation differences in shell growth rates and
hence, affect the degree of shell inflation.

For example, specimens of Corbicula fluminea in the
collections of the Delaware Museum of Natural History (DMNH
110469) and Texas Christian University (TCU 6087) from un-
named irrigation ditches at Montezuma’s Well National Monu-
ment, Yavapai County, Arizona, demonstrate extreme lateral
compression of the valves. Yet, this condition is regarded as
an expression of response to environmental conditions rather
than indicative of another species (Britton and Morton, 1986).

Prezant and Chalermwat found that shell microstructure (1983)
and internal shell color changes (1983, 1984) could be induced
in C. fluminea by alteration of the thermal and trophic regimes.
While color change could be induced only from purple to
white, Prezant and Chalermwat speculated that many of the
morphological differences seen in North American corbiculids
could be a reflection of microhabitat rather than species-
specific differences.

Britton and Morton (1986) and Morton (1987) further
noted the polymorphism among populations of Corbicula
fluminea in North America and Hong Kong, respectively.
These studies report differences in shell morphology and color
on the basis of sex as well as differences in water quality. Mor-
ton (1987) particularly noted that pH, dissolved oxygen and
carbon dioxide, and potassium were highly correlated with
morphological differences in C. fluminea of Hong Kong. Brit-
ton and Morton (1986) found that shell characters were
unreliable to differentiate between the color morphs of C.
fluminea North America. On the basis of this and other
ecological data, they concluded that there was only one
species of Corbicula in North America. These considerations
are not addressed in any detail by Soviet malacologists with
respect to species determination. As yet, no malacologist in
the U.S.S.R. has attempted to resolve systematic problems
within the genus Corbicula using electrophoretic techniques.

ACKNOWLEDGMENTS

| wish to thank Drs. Yaroslav |. Starobogatov and Alexander
Golikov, Zoological Institute, Academy of Sciences of the U.S.S.R.,
Leningrad, for their many kindnesses, discussions, and assistance
during my stay at the institute. Thanks are also due Dr. Z. |. Izzatullaev,
Institute of Zoology and Parasitology, Academy of Sciences,
Dushanbe, Tadzhik S.S.R., for his discussions on the corbiculids of
Central Asia. Two anonymous reviewers made helpful suggestions
that improved the paper. Finally, | wish to thank Diana B. Bieliauskas
and the staff of the National Academy of Sciences, Advisory Com-
mittee on the U.S.S.R. and Eastern Europe, for their assistance in
arranging my studies in the Soviet Union.

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Date of manuscript acceptance: 3 February 1989

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00 .00 615.50 1.26

00 .00 250.00 51

00 .00 62.50 13
$3,425.18 87.14 $21,856.48 44.60
$3,930.68 100.00 $49,004.28 100.03

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Current-Period
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$ 15.00 03
196.01 40
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369.93 75
805.33 1.64
113.50 .23
379.66 77
63.87 13
185.10 38
560.00 1.14
2,265.00 462
49.16 10
2,000.00 4.08
750.00 1.53
231.00 47
22.66 05
13.00 03
431.70 88
14,350.54 29.28
52 02

135.79 .28
$42,120.17 85.93
$ 6884.11 14.10

55TH ANNUAL MEETING
THE AMERICAN MALACOLOGICAL UNION
LOS ANGELES, CALIFORNIA
JUNE 25 - 30, 1989

The 55th annual meeting of the American Malacological Union will be a combined
meeting with the Western Society of Malacologists, held June 25-30, 1989, in Los
Angeles, California, at the Davidson Conference Center of the University of Southern
California. Facilities at the Los Angeles County Museum of Natural History will also
be used for some of the events. There are to be three choices for housing, the
University Hilton, the Vagabond Motel, and the University dormitories, all very close
to the Davidson Conference Center. In addition to the proximity of beaches and moun-
tains, the Los Angeles area has a delightful summer climate in late June, almost never
too hot or humid.

Three symposia are planned:

BIOLOGY OF PELAGIC GASTROPODS
(Organized by Dr. Roger Seapy)

SYSTEMATICS AND EVOLUTION OF WESTERN NORTH AMERICAN LAND MOLLUSKS
(Organized by Drs. F. G. Hochberg and Barry Roth)

BIOLOGY OF SCAPHOPODS
(Organized by Dr. Ronald L. Shimek)

In addition to the symposia, contributed papers, and poster presentations, scheduled events will
include field trips, an outdoor barbecue, and a banquet.

For further information please contact:
James H. McLean
President, AMU
Los Angeles County Museum of Natural History
900 Exposition Boulevard
Los Angeles, California 90007, U.S.A.

(213) 744-3377

In Errata: Volume 6, No. 2, page 219: Under dates of publication and key, change ‘‘Volume 6, No. 2: July 1988 [6(2)]”’
to read ‘‘Volume 6, No. 2: October 1988 [6(2)].”’

89

SPECIAL PUBLICATIONS OF THE
AMERICAN MALACOLOGICAL BULLETIN

The Special Publication Series of the American Malacological Bulletin was begun to
disseminate collected sets of papers with similar or related themes in a single volume.
To date, three such issues have been published, each the result of a special convened
symposium. The three Special Editions are PERSPECTIVES IN MALACOLOGY, PRO-
CEEDINGS OF THE SECOND INTERNATIONAL CORBICULA SYMPOSIUM, and PRO-
CEEDINGS OF THE SYMPOSIUM ON ENTRAINMENT OF LARVAL OYSTERS. Additional
Special Editions are planned for the near future.

PERSPECTIVES IN MALACOLOGY (Sp. Ed. #1, July, 1985) offers a wide range of papers

dealing with molluscan biology of interest to professionals and amateurs alike. These

papers were presented as part of asymposium held in honor of Professor M. R. Carriker
at the time of his retirement and highlight a variety of recent advances in numerous facets of the study of molluscs. PERSPEC-
TIVES IN MALACOLOGY offers insight into some of the frontiers of molluscan biology ranging from deep-sea hydrothermal
vent malacofauna to chemical ecology of oyster drills.

The PROCEEDINGS OF THE SECOND INTERNATIONAL CORBICULA SYMPOSIUM (Sp. Ed. #2, June 1986) contains
numerous papers on this exotic bivalve that has become a significant “‘pest’’ organism of several power plants and other
industries using cooling waters. The proliferation, spread, functional biology, attempts at industrial control, taxonomy,
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The third special edition of the American Malacological Bulletin, PROCEEDINGS OF THE SYMPOSIUM ON THE ENTRAIN-
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VOLUME 7 1990 NUMBER 2

CONTENTS

Genetic variation in Neotricula aperta, the intermediate snail host of
Schistosoma mekongi: allozyme differences reveal a group of
sibling species. KATHARINE C. STAUB, DAVID S. WOODRUFF,.
E. SUCHART UPATHAM and VITHOON VIYANANT .................... 0.0.00. e eee eee 93

Cellular DNA contents of the freshwater snail genus Semisulcospira
(Mesogastropoda: Pleuroceridae) and some cytotaxonomical remarks.
HIROSHI K. NAKAMURA and YOSHIO OJIMA ............. 00.0.0. 0. ee 105

Use of shell morphometric data to aid ciassification of Pisidium
(Bivalvia: Sphaeriidae). BRUCE W. KILGOUR, DENIS H. LYNN
eMC EENE Lg MIG INES st ik, Oh ec Ride baste oi bre N ee kd ee ote ewe cael 109

Polymorphism for shell color in the Atlantic Bay Scallop Argopecten
irradians irradians (Lamarck) (Mollusca:Bivalvia) on Martha’s
Vineyard Island. J. A. ELEK and S. L. ADAMKEWICZ ................. 0.20.00. eee AIT

Prehistoric freshwater mussel (naiad) assemblages from southwestern
ona warie Sten Mein ye titer ees oo er Ra ee ak te a Py. le BR ee ae 127

Research Note: Rectification of the nomenclature of certain
species of Triculine snails transmitting Paragonimus
and Schistosoma in China. LIU YUE LING and
SE ORGEENT YORI Sia Bee eee AE harks ce Ph ME A Bee ee Ne ee 131

SYMPOSIUM ON THE BIOLOGY OF THE SCAPHOPODA

Functional morphology of the perianal sinus and pericardium of
Dentalium rectius (Mollusca: Scaphopoda) with a

reinterpretation of the scaphopod heart. PATRICK D. REYNOLDS ....................... 137
Diet and habitat utilization in a northeastern Pacific Ocean scaphopod

BopempiageRONAL Daly SHIMEK Ss es A Bis ea Scie ee Gn de oe ee ak 147
PeipenCIAE EOS eis ths Aroha cS) 2 LNA da teeteteete ae ae ee eee Game Se 171

PAMOUNCEINE MSs eae eras anh sRenA ome nts owen ees a ee a at eee. 173

LIBRARIES

INGO ROR VOIUIMES Fo us See ts DA ek eS PTR, NN,

AMERICAN MALACOLOGICAL BULLETIN

EDITOR-IN-CHIEF

ROBERT S. PREZANT
Department of Biology
Indiana University of Pennsylvania
Indiana, Pennsylvania 15705

MELBOURNE R. CARRIKER
College of Marine Studies

University of Delaware
Lewes, Delaware 19958

GEORGE M. DAVIS

Department of Malacology

The Academy of Natural Sciences
Philadelphia, Pennsylvania 19103

R. TUCKER ABBOTT
American Malacologists, Inc.
Melbourne, Florida, U.S.A.

JOHN A. ALLEN
Marine Biological Station
Millport, United Kingdom

JOHN M. ARNOLD
University of Hawaii
Honolulu, Hawaii, U.S.A.

JOSEPH C. BRITTON
Texas Christian University
Fort Worth, Texas, U.S.A.

JOHN B. BURCH
University of Michigan
Ann Arbor, Michigan, U.S.A.

EDWIN W. CAKE, JR.
Gulf Coast Research Laboratory
Ocean Springs, Mississippi, U.S.A.

PETER CALOW
University of Sheffield
Sheffield, United Kingdom

BOARD OF EDITORS

MANAGING EDITOR

ASSOCIATE EDITORS

RONALD B. TOLL
Department of Biology
University of the South

Sewanee, Tennessee 37375

W. D. RUSSELL-HUNTER
Department of Biology

Syracuse University
Syracuse, New York 13210

ROGER HANLON

Ex Officio

Marine Biomedical Institute
University of Texas
Galveston, Texas 77550

BOARD OF REVIEWERS

JOSEPH G. CARTER
University of North Carolina
Chapel Hill, North Carolina, U.S.A.

ARTHUR H. CLARKE
Ecosearch, Inc.
Portland, Texas, U.S.A.

CLEMENT L. COUNTS, III
University of Maryland
Princess Anne, Maryland, U.S.A.

THOMAS DIETZ
Louisiana State University
Baton Rouge, Louisiana, U.S.A.

WILLIAM K. EMERSON
American Museum of Natural History
New York, New York, U.S.A.

DOROTHEA FRANZEN
Illinois Wesleyan University
Bloomington, Illinois, U.S.A.

VERA FRETTER

University of Reading
Berkshire, United Kingdom

ISSN 0740-2783

THOMAS R. WALLER
Department of Paleobiology
Smithsonian Institution
Washington, D. C. 20560

JOSEPH HELLER
Hebrew University of Jerusalem
Jerusalem, Israel

ROBERT E. HILLMAN
Battelle, New England
Duxbury, Massachusetts, U.S.A.

K. ELAINE HOAGLAND
Association of Systematics Collections
Washington, D.C., U.S.A.

RICHARD S. HOUBRICK
U.S. National Museum
Washington, D.C., U.S.A.

VICTOR S. KENNEDY
University of Maryland
Cambridge, Maryland, U.S.A.

ALAN J. KOHN
University of Washington
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University of Arkansas
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JOHN N. KRAEUTER
Baltimore Gas and Electric
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ALAN M. KUZIRIAN

NINCDS-NIH at the

Marine Biological Laboratory
Woods Hole, Massachusetts, U.S.A.

RICHARD A. LUTZ
Rutgers University
Piscataway, New Jersey, U.S.A.

EMILE A. MALEK
Tulane University
New Orleans, Louisiana, U.S.A.

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University of Southern Maine
Portland, Maine, U.S.A.

JAMES H. McLEAN
Los Angeles County Museum
Los Angeles, California, U.S.A.

ROBERT F. MCMAHON
University of Texas
Arlington, Texas, U.S.A.

ROBERT W. MENZEL
Florida State University
Tallahassee, Florida, U.S.A.

ANDREW C. MILLER
Waterways Experiment Station
Vicksburg, Mississippi, U.S.A.

BRIAN MORTON
University of Hong Kong
Hong Kong

JAMES J. MURRAY, JR.
University of Virginia
Charlottesville, Virginia, U.S.A.

RICHARD NEVES

Virginia Polytechnic Institute
and State University

Blacksburg, Virginia, U.S.A.

JAMES W. NYBAKKEN
Moss Landing Marine Laboratories
Moss Landing, California, U.S.A.

WINSTON F. PONDER
Australian Museum
Sydney, Australia

CLYDE F. E. ROPER
U.S. National Museum
Washington, D.C., U.S.A.

NORMAN W. RUNHAM
University College of North Wales
Bangor, United Kingdom

AMELIE SCHELTEMA
Woods Hole Oceanographic Institution
Woods Hole, Massachusetts, U.S.A.

ALAN SOLEM
Field Museum of Natural History
Chicago, Illinois, U.S.A.

DAVID H. STANSBERY
Ohio State University
Columbus, Ohio, U.S.A.

FRED G. THOMPSON
University of Florida
Gainesville, Florida, U.S.A.

THOMAS E. THOMPSON
University of Bristol
Bristol, United Kingdom

NORMITSU WATABE
University of South Carolina
Columbia, South Carolina, U.S.A.

KARL M. WILBUR
Duke University
Durham, North Carolina, U.S.A.

Cover. This illustration of Bathyliotina glassi was used on the logo of the 1989 annual meeting of the American Malacological Union. Papers
resulting from the symposium entitled ‘‘Biology of Scaphopods’,, held at that meeting, can be found in this volume, beginning on page 137.

THE AMERICAN MALACOLOGICAL BULLETIN is the official journal publication of the American Malacological Union.

AMER. MALAC. BULL. 7(2)
February 1990

GENETIC VARIATION IN NEOTRICULA APERTA, THE INTERMEDIATE
SNAIL HOST OF SCHISTOSOMA MEKONGI: ALLOZYME
DIFFERENCES REVEAL A GROUP OF SIBLING SPECIES

KATHARINE C. STAUB
DAVID S. WOODRUFF
DEPARTMENT OF BIOLOGY (C-016) AND
CENTER FOR MOLECULAR GENETICS
UNIVERSITY OF CALIFORNIA, SAN DIEGO
LA JOLLA, CALIFORNIA 92093, U.S.A.

E. SUCHART UPATHAM
VITHOON VIYANANT
CENTER FOR APPLIED MALACOLOGY AND ENTOMOLOGY
DEPARTMENT OF BIOLOGY, FACULTY OF SCIENCE
MAHIDOL UNIVERSITY
RAMA VI ROAD, BANGKOK 10400, THAILAND

ABSTRACT

Neotricula aperta (Temcharoen) is a highly variable pomatiopsid gastropod found in the Mekong
River and its tributaries in Thailand. Three races and two other variant phenotypes have been described,
originally as Tricula aperta. Samples of the sympatric alpha and gamma races from the Mekong River
and of the beta race from the Mun River were characterized at 16 allozyme loci. Highly significant
heterozygote deficiencies and differences in allele frequencies between males and females were ap-
parent at many polymorphic loci in each of the three samples. The observed heterozygote deficien-
cies and sexual differences were artifacts produced by the presence of cryptic taxa in each original
racial sample. In the Mun River beta race sample, we found two sibling species separated by a signifi-
cant multilocus genetic distance (D = 0.22). In the Mekong River, the snails representing the alpha
and gamma races were found to be referable to two other well differentiated sibling species (D = 0.34),
both of which have individuals of ‘‘alpha’’ and ‘‘gamma’’ morphotypes. The Mekong River species
pair are very well differentiated from the Mun River species pair (D = 0.74). Formal taxonomic revision
of the N. aperta sibling species complex is postponed until topotypic material (N. aperta gamma race)
from Laos can be examined. As only the ‘‘gamma race’ had been shown to transmit Schistosoma
mekongi naturally, it remains to be established which of the newly recognized species are
epidemiologically significant.

The major Late Tertiary radiation of Triculinae (Pro-
sobranchia: Rissoacea: Pomatiopsidae) in Southern China
and Southeast Asia has resulted in more than 12 genera and
120 species of small freshwater snails (Davis, 1979, pers.
comm., 1986; Kang, 1983, 1984a, b, 1986; Liu et a/., 1983).
Neotricula aperta (Temcharoen) is the best known member
of this extraordinary radiation as it is the intermediate host
for the human blood fluke, Schistosoma mekongi Voge,
Bruckner and Bruce. In this paper we will present evidence,
based on multilocus allozyme variation, suggesting that

N. aperta actually comprises a group of at least four sibling
species.

The species was first described as Lithoglyphopsis
aperta by Temcharoen (1971) and subsequently placed in the
genus Tricula by Davis (1979), and Neotricula by Davis et al.
(1986). These are small (2-4 mm shell length), dioecious,
aquatic prosobranch snails. In their monograph, Davis et al.
(1976) described three races of this species in Thailand and
Laos on the basis of shell size, shape, and microsculpture,
mantle pigmentation, developmental rates, radular traits,

American Malacological Bulletin, Vol. 7(2) (1990):93-103

93

94 AMER. MALAC. BULL. 7(2) (1990)

features of male reproductive anatomy, habitat and distribu-
tion. Kitikoon et al. (1981) described the last two traits of the
three races in more detail. The alpha and gamma races are
found, frequently together, along 300 km of the Mekong River.
The beta race is found only in the Mun River (alternatively
transliterated as Mool), a tributary of the Mekong in northeast
Thailand. Shell size, shell shape, and mantle pigmentation
have been the diagnostic characters used in field identifica-
tion for the sympatric alpha and gamma races. Gamma race
snails typically have four large, distinctive pigment spots on
their mantles that are absent in alpha and beta race snails.
Gamma race snails are also often smaller than sympatric
alpha race snails; beta race snails are intermediate in size.
In the Mekong River, alpha and gamma race snails occupy
the same range of benthic microhabitats: from near shore to
river center and also in seasonal pools on exposed rock
islands. Beta race snails occur in and near rapids in the Mun
River. Snails of all races are found on solid substrata (rocks
and sticks) and never on sand, mud or algal strands.

Davis et al. (1976) discussed the possibility that the
three races could be reproductively isolated from one another.
They suggested that the beta race, with its allopatric distribu-
tion and pronounced microhabitat preferences, could be
specifically distinct from the Mekong River races. They further
speculated that differential rates of growth and maturation
could act to isolate the sympatric alpha and gamma races
reproductively, and that these two taxa also could have
reached full species rank. They concluded, however, that their
evidence for significant intraracial variation and for racial in-
termediacy did not support such conclusions. They argued
that the known differences in the reproductive organs, shell
size and sculpture, pigmentation, and radular formulae could
simply reflect differences in ontogeny and ecology rather than
genetically-based evolutionary divergence. They found ap-
parent hybrids (snails with irregular pigment patterns and shell
size and shape intermediate between the alpha and gamma
races) at one locality and noted that the alleged anatomical
differences between these races were somewhat artificial.
Similarly, microhabitat preferences overlap broadly (Kitikoon
et al., 1981). Thus, no formal subspecific nomenclature was
proposed to partition the variation recognized in this species
from the outset.

This view of Neotricula aperta as a highly variable
species with recognizable ecophenotypic races was subse-
quently challenged by Kitikoon’s reports (1981a, b, 1982a, b;
Kitikoon et al., 1981) on additional phenotypic variation,
chromosome numbers and karyotypes, isoenzyme patterns,
and parasite compatibility. Kitikoon (1982b:55) suggested that
‘the so-called alpha, beta, and gamma ‘“‘races”’ of T. aperta
are at least different subspecies and may well be different
species. Our quantitative population study of allozyme varia-
tion in this taxon supports the latter conclusion, but in a man-
ner completely unanticipated in our preliminary report
(Woodruff et a/., 1986b).

MATERIALS AND METHODS

Using distributional, ecological, and morphological

criteria to identify snails in the field, samples of Neotricula
aperta alpha, beta, and gamma races were collected in north-
eastern Thailand in May 1984. N. aperta alpha race and
gamma race snails were taken from the Mekong River near
Ban Bungkhong in the Khemarat District of Ubon Ratchathani
Province. Alpha snails were found in pools on a rock island
and gamma race snails were taken nearby from rocks
cropping out in the main river channel where the water was
deeper and the current swifter. In both cases, however, water
depths in May were less than 1.0 m. N. aperta beta race snails
were collected in the Mun River near Khaeng Khao in the
Phibun Mangsuhan District of Ubon Ratchathani Province.
This site is midway between the town of Ubon and the Mun’s
juncture with the Mekong and about 100 km directly south
of the alpha-gamma collection site. Snails were taken from
rocks in fast flowing water less than one meter deep. In every
case, sampling was conducted along less than 10 m of river
bottom. Racial identities were confirmed and snails were
sexed under a binocular microscope in Bangkok soon after
collection. Snails were then frozen at -70°C until elec-
trophoresis was carried out at the University of California, San
Diego in 1985. Voucher specimens were deposited in the
museum at the Center for Applied Malacology, Mahidol
University.

The electrophoretic techniques used are described in
general terms elsewhere (Mulvey and Vrijenhoek, 1981;
Woodruff et a/., 1988). Individual snails were quickly
homogenized in less than 0.1 ml (2-3 drops from a standard
Pasteur pipette) of grinding solution (0.01 M Tris, 0.001 M EDTA,
0.05 mM NADP, pH 7.0) with a glass rod. The homogenate
was centrifuged at 10,000 g for 2 min in a Fisher 235A micro-
centrifuge, and the supernatant was absorbed onto 3 x 9mm
tabs of Whatman No. 3 chromatography paper which were
then inserted into cold 12% Sigma® starch gels (one tab per
snail per gel). Electrophoresis was carried out using four dif-
ferent buffer systems at 4°C for 15-18 hrs (Table 1). A
bromophenol blue marker dye migrated 100-125 mm anodal-
ly during this time except in the case of buffer system Tris-
Citrate pH 6.8 in which the marker migrated 70-100 mm.
Following electrophoresis, 4-5 slices were cut from each gel
and each slice was stained for a specific enzyme following
standard methods (Shaw and Prasad, 1970; Harris and
Hopkinson, 1978). The esterase substrate was alpha-napthyl
acetate; the peptidase substrate was leucyl-alanine.

Electrophoretic conditions for the resolution of 12
enzymes coding for the 16 allozyme loci reported here are
described in Table 1. These enzymes were selected on the
basis of their interpretable electromorphs from about 30 en-
zymes tested under ten different electrolyte and pH condi-
tions and are associated with a variety of metabolic pathways.
Snails from different samples were run on each gel to facilitate
commparisons; isozymes were numbered and allozymes
assigned mobility values relative to the common electromorph
in Neotricula aperta alpha race. In Table 6, alleles are listed
in order of their decreasing anodal mobilities; cathodal mobili-
ty is indicated by a negative value. For each locus, relative
mobilities are those for the first buffer system reported in
Table 1. These are reported to two decimal places only where

STAUB ET AL.: GENETIC VARIATION IN NEOTRICULA APERTA 95

Table 1. Electrophoretic buffers used for resolution of proteins in Neotricula aperta.

Enzyme (E. C. #)

Abbreviation Buffer”

Acid phosphatase (3.1.3.2) ACP TC 68
Aspartate aminotransferase (2.6.1.1.) AAT TBE 80
Esterase (3.1.1.1) EST-1 TBE 80
EST-2 TBE 80
EST-3 TBE 80
Glyceraldehyde-3-phosphate dehydrogenase (1.2.1.12) GAP AP 6.0, TC 68
Glycerol-3-phosphate dehydrogenase (1.1.1.8) GPDH AP 6.0, TC 68
Glucose phosphate isomerase (5.3.1.9) GPI TC 6.0, TC 68
Isocitrate dehydrogenase (1.1.1.42) IDH TC 68
Leucine aminopeptidase (3.4.11) LAP (PEP-2) TC 6.0, TBE 80
Malate dehydrogenase (1.1.1.37) MDH TC 6.0
Peptidase (3.4.-) PEP-3 TBE 8.0
PEP-4 TBE 80
6-Phosphogluconate dehydrogenase (1.1.1.44) PGD AP 6.0
Phosphoglucomutase (5.4.2.2) PGM-1 TC 6.0
PGM-2 AP 6.0

*AP 6.0: 0.04 M citrate adjusted with N-(3-aminopropyl)-morpholine to pH 6.0; diluted 1:19 for gels and
undiluted for electrodes (16 hr., 80 v). TBE 8.0: 0.5 M Tris, 0.65 M borate, 0.02 M EDTA, adjusted to
pH 80; diluted 1:9 for gels and undiluted for electrodes (16 hr., 100 v). TC 6.0: 0.378 M Tris, 0.165 M
citrate, adjusted to pH 6.0; 13.5 ml diluted to 400 ml for gel and undiluted for electrodes (16 hr., 60 v).
TC 68: 0.188 M Tris, 0.065 M citrate, adjusted to pH 68: diluted 1:9 for gels and 1:5 for electrodes

(16 hr., 150 v).

they cannot be distinguished by a single decimal place ap-
proximation. Commonly used enzyme abbreviations are
typeset in capital letters to indicate the protein and in italics
to indicate the presumed locus.

The mean number of alleles per locus (A), the propor-
tions of loci polymorphic (a locus was considered polymor-
phic if more than one allele was detected,) P. and the mean
individual heterozygosity (by direct count) (H), were calculated
for each sample. Allozyme frequencies for the polymorphic
loci were tested for their agreement with Hardy-Weinberg ex-
pectations for a panmictic population by X2- test where ap-
propriate and by the Fisher exact test. Allozyme frequency
differences between sexes were also tested for significance
by X2- and G-tests. Genetic distance coefficients (D) (Nei,
1978; standard error after Nei et a/., 1985) and genetic similari-
ty coefficients (S) (Rogers, 1972) were calculated and clustered
by the UPGMA algorithm. X2- and G-tests were performed
with software accompanying Sokal and Rohlf (1981); other
analyses were performed with the BIOSYS-1 computer pro-
gram (Swofford and Selander, 1981).

The above analyses were first performed on the original
“racial’’ samples with sexes pooled and then with sexes
separated. As the original samples were found to be highly
heterogeneous, it became necessary to repeat the analyses
with the snails from each sample site resorted according to
individual genotype. The sorting procedure, based on three
or more diagnostic loci, is described below.

RESULTS

VARIATION IN THE THREE ORIGINAL SAMPLES
SORTED INTO ALPHA, BETA AND
GAMMA RACES

For reasons that will become clear below, the allele fre-

quencies at all 16 presumptive loci are not reported here.
These data on variation in the original alpha, gamma and beta
“racial’”’ samples are presented elsewhere (Staub, 1988).
Our preliminary analyses showed Neotricula aperta
alpha race and gamma race samples were virtually in-
distinguishable at all loci (D = 0.01 + 0.02). In contrast, the
mean genetic distance value between beta race and the two
Mekong River races was unexpectedly large (D = 0.66 +
0.24). However, panmixia is an important assumption of Nei’s
genetic distance statistics and significant departures from
Hardy-Weinberg expectations for panmixia were detected in
62% on the polymorphic loci in these samples (Table 2). In
all 18 cases, there was a deficiency of heterozygotes and all
but two tests were significant at the 1% (0 <0.01) level.
The ratio of females to males in the original racial
samples was 32:36 for alpha race, 34:43 for beta race, and
32:34 for gamma race. In each original sample, the pattern
of loci with heterozygote deficiencies was essentially the same
within each sex as it was with the sexes pooled. Neither males

Table 2. Number of loci showing a significant deficiency of
heterozygotes, as a fraction of the total number of polymorphic loci,
before and after resorting Neotricula aperta racial samples by three-
locus genotype.**

Original Samples Resorted Samples

alpha race 6(6)*/10 Mekong River taxon 1 2(0)/8
gamma race 7(6)/11 Mekong River taxon 2 2(2)/10
beta race 5(4)/8 Mun River taxon 1 0(0)/6

Mun River taxon 2 4(2)/6

“Number of Fisher exact tests significant at 0.01 < p <0.05 and at
p <0.01 (in parentheses).
*“See text and Tables 4 and 5 for full explanation.

96 AMER. MALAC. BULL. 7(2) (1990)

nor females contributed more to any sample’s overall defi-
ciencies and no single-locus genotype appeared to be sex-
linked for any sample. However, allele frequencies were
notably different between the sexes at many loci. This too was
unexpected as males and females allegedly represent a ran-
dom sample of each population and sex-linked allozymes are
rare (Richardson et al., 1986). Tests of sample independence
between male and female subsets revealed significant (p
<0.05) differences in each original sample (Table 3).

Although the initial analysis suggested that the Gap’-°
and Gap'-4 alleles were equally abundant in the alpha and
gamma races (Staub, 1988), no heterozygotes were observed
among 118 animals. Likewise, no Pep-31-2/1-0 heterozygotes
were observed among 120 animals. Concordance by specific
genotype between these two loci and a third with a marked
deficiency of heterozygotes, Gpi (N = 126) was 100%
(Table 4). Similarly, no heterozygotes were observed at three
polymorphic loci in the beta race sample: Lap (N = 59), Mdh
(N = 65), and Pep-3 (N = 48) and the concordance by
genotype between these three loci is also nearly complete
(Table 5).

The unexpected differences between sexes and these
striking associations among alleles at loci with no heterzygotes
suggested that the original sample sorting had been insen-

Table 3. Number of loci showing a significant difference in allele fre-
quencies between males and females, as a fraction of the total number
of polymorphic loci, before and after resorting Neotricula aperta
“racial’’ samples by three-locus genotype.**

Original Samples Resorted Samples

alpha race 6(2)*/10 Mekong River taxon 1 1(0)/8
gamma race 5(4)/11 Mekong River taxon 2 1(0)/10
beta race 2(1)/8 Mun River taxon 1 eal

Mun River taxon 2 2(0)/6

*Number of Fisher exact tests significant at 0.01 <p <0.05 and at
p<0.01 (in parentheses).
**See text and Tables 4 and 5 for full explanation.
***Sample size too small to analyze.

sitive to the genetic heterogeneity present at each locality.
The snails from each original collecting locality were accord-
ingly resorted by three-locus genotype, as identified in Tables
4 and 5, and the analyses were repeated. The original racial
designations were abandoned.

VARIATION IN MEKONG RIVER AND MUN RIVER
SAMPLES FOLLOWING REASSORTMENT
BY INDIVIDUAL MULTILOCUS GENOTYPE

The alpha race and gamma race samples were pooled
as a Mekong River sample within which we discovered two
genetically defined groups of individuals, hereafter called
Mekong River taxon 1 and Mekong River taxon 2. Each of
these newly recognized taxa includes snails previously
referred to both alpha and gamma races. The ratio of alpha
race to gamma race snails was 30:34 and 38:32 for taxon 1
and taxon 2 respectively; the ratio of female to male snails
in the new taxa was 32:32 and 32:38, respectively.

Similarly, the snails in the Mun River sample original-
ly referred to the beta race are hereafter assigned to two
genetically defined groups: Mun River taxon 1 and Mun River
taxon 2. The ratio of female to male snails was 3:8 and 31:31
for taxon 1 and taxon 2, respectively. Three snails had ap-
parently intermediate genotypes but were assigned to taxon
2 on the basis of variation at Mdh and Pep-3 only because
the Lap'-°5 and Lap'-° electromorphs are too similar for reliable
discrimination of all individuals. Four snails could not be
assigned because they did not show activity for any of the
three diagnostic enzymes.

Allele frequencies for the newly recognized taxa are
displayed in Table 6 along with summary genetic variability
statistics. The number of polymorphic loci is eight in Mekong
River taxon 1 and ten in Mekong River taxon 2. There are
striking differences in allele frequencies between these two
taxa at Est-7, Pep-4, Pgm-1, and Pgm-1, as well as at the three
loci (Lap, Mdh, and Pep-3) on which the reassortment was
based -- in other words, at virtually all the polymorphic loci.
This observation is quantified by the significant genetic
distance value for the two taxa, D=0.34 + 0.16.

Table 4. Genotypes at three electrophoretic loci for 134 snails originally referred to Neotricula aperta alpha race
and Neotricula aperta gamma race, showing Mekong River taxon to which snails of each genotype were assigned.

Locus No. snails*
Mekong River (118)** (126) (120)
taxon Gap Gpi Pep-3 N3 No N,
1 1.4/1.4 1.0/1.0 1.2/1.2 42 10 2
1 1.4/1.4 1.0/1.0 1.2/1.11 4 3 0
1 1.4/1.4 1.0/1.0 1.19/1.11 3 0 0
2 1.0/1.0 2.0/2.0 1.0 /1.0 27 16 0
2 1.0/1.0 2.0/2.0 1.11/1.0 2 2 0
2 1.0/1.0 2.0/1.0 1.0/1.0 18 3 0
2 1.0/1.0 1.0/1.0 1.11/1.0 1 0 0
2 1.0/1.0 1.0/1.0 1.0/1.0 1 0 0

*N3 = number of individuals scored at all three loci; No = number scored at two of the three loci; N; = number

scored at one locus.

*“Total number of individuals scored at this locus shown in parentheses.

STAUB ET AL.: GENETIC VARIATION IN NEOTRICULA APERTA 97

Among the Mun River taxa, formerly lumped as beta
race, the number of polymorphic loci is five in Mun River tax-
on 1 and six in Mun River taxon 2 (Table 6). There are notable
differences in allele frequencies between these two taxa at
Pgd and Est-3 in addition to the three loci (Lap, Mdh, and
Pep-3) used to resort the original sample -- again, at virtually
all the polymorphic loci. The genetic distance value for the
two Mun River taxa is D = 0.22 * 0.12.

The newly recognized Mekong River taxa show
markedly fewer heterozygote deficiencies than the original
alpha race and gamma race samples (Table 2). There are
residual deficiencies (0 <0.05) at Lap and Pep-3 in Mekong
River taxon 1 and at Acp and Pep-4 in Mekong River taxon
2; two of these four positive tests are significant at the 1%
level, those for Acp and Pep-4 in Mekong River taxon 2. In
the Mun River taxa, the number of loci showing a deficiency
of heterozygotes is also reduced compared to the original beta
race sample (Table 2). The sample of Mun River taxon 1 is
too small to analyze. Deficiencies remain at Est-7, Lap, Pgd,
and Pgm-7 in Mun River taxon 2; two of the four positive tests
are significant at the 1% level, those for Est-7 and Lap.

Differences in allele frequencies between the sexes
were eliminated by the reassortment of alpha and gamma
races into Mekong River taxon 1 and 2 (Table 3). A difference
(0.01 <p<0.05) remains at the Acp locus for both Mekong
River taxa. The effect of the beta race reassortment on sex
differences in allele frequencies cannot be established for
Mun River taxon 1 as the sample is too small to analyze (N
= 11). In Mun River taxon 2, two loci (Acp and Pgd) show
minor (and statistically insignificant) sex-related differences.
In each of the four newly recognized taxa, the pattern of loci
with heterozygote deficiencies is essentially the same within
each sex as it had been for the sexes pooled. Neither males
nor females contributed more to any sample’s overall defi-
ciencies and no single-locus genotype appeared to be sex-
linked for any sample.

The genetic distance value between the Mekong River
taxa and the Mun River taxa is very significant (D=074 +
0.26), as depicted in the phenogram (Fig. 1). Table 7 displays
genetic distance and identity values for each of the six pair-
wise comparisons for the four taxa.

DISCUSSION

HETEROZYGOTE DEFICIENCIES AND THEIR
INTERPRETATION

The validity of our conclusion depends on our inter-
pretation of the massive heterozygote deficiencies as evidence
for a type of sampling error commonly referred to as the
Wahlund effect. However, a discussion of other reasons for
heterozygote deficiencies is appropriate here as there are
several other possibilities that demand critical consideration.
Heterozygote deficiencies across most or all polymorphic en-
zyme loci have been reported for other mollusc populations,
particularly among marine species (Johnson and Black, 1984;
McMeekin, 1985; Singh and Green, 1984; Woodruff et al.,
1986a; Zouros and Foltz, 1984) and a number of potential

causes have been considered. These can be roughly
classified into two groups: first, true deficiencies of
heterozygotes in natural populations and, second, apparent
deficiencies due to sampling error or other experimental er-
ror. A true heterozygote deficiency at an enzyme locus could
be due to (1) location of the locus on a sex chromosome
(Zouros et al., 1980), (2) complete or partial inbreeding
(Hedrick and Cockerham, 1986), or (3) selection against
heterozygotes, for instance, at a particular developmental
stage (Singh and Green, 1984). There are, in addition, a
number of ways in which heterozygote deficiency at a locus
could be apparent, but not real, due to (4) scoring bias for
homozygotes (Ayala et a/., 1973), (5) presence of one or more
null alleles (Zouros et al., 1980), (6) biased sampling of
homozygotes, or (7) the so-called Wahlund effect (Singh and
Green, 1984). Biased sampling of homozygotes could be due
to (6a) genetic patchiness across a population’s habitat at the
time of collection, referred to as population subdivision by
Zouros et al. (1980), or (6b) differential survival of homozygotes
following collection. The Wahlund effect is the result of mix-
ing representatives of two (or more) independent gene pools
with differing allele frequencies in the same sample. It could
be due to (7a) the existence of cryptic (sibling) taxa or (7b)
error in field identification and taxonomic separation among
already recognized taxa showing similar features. We con-
sider each of these hypotheses in turn.

1. Real heterozygote deficiencies due to chromosomal con-
straints. The mechanism of chromosomal determination of sex
in Neotricula aperta is not known but was speculated by
Kitikoon (1982a) to be an XO-male/XX-female mechanism,
based on chromosome pairing data. Such a system would
result in an absence of heterozygotes in males at any X-linked
locus. We did not find sex-linkage at any locus, either before
or after the reassortment (29 and 24 tests, respectively).
Moreover, such a system (or any other) does not predict the
generalized (multilocus) strong genotypic disequilibria shown
by our data.

2. Real heterozygote deficiencies due to inbreeding.
Although self-fertilization is not possible in dioecious triculines,
it is possible that partial inbreeding could be contributing to
increased homozygosity in these snails. Neotricula aperta
snails have low vagility and are substrate limited (Davis et al.,
1976); they are found packed tightly on solid substrata, often
rocks, presumably where the eggs from which they hatched
were deposited, and it is possible that sib matings occur with
high frequency. The snails fit the profile of r-selected colonists
whose patchily distributed habitats are changed annually
when the monsoon flood raises river levels dramatically (up
to 15 m in the Mekong) (Davis et a/., 1976). Female survivor-
ship from the preflooding time of copulation to the postflooding
time of egg deposition is low and founder effects could further
increase the likelihood of sib matings. As inbreeding affects
all loci in a uniform way, the generalized disequilibria shown
by our data are consistent with this hypothesis. However, the
complete absence of intermediates between the two
genetically-defined Mekong River taxa negates this
hypothesis.

3. Real heterozygote deficiencies due to natural selection.

98 AMER. MALAC. BULL. 7(2) (1990)

Table 5. Genotypes at three electrophoretic loci for 74 snails originally referred to Neotricula aperta beta race

’

showing Mun River taxon to which snails of each genotype were assigned.

Locus No. snails*
Mun River (59)** (66) (46)
taxon Lap Mdh Pep-3 N3 No N,
1 1.0/1.0 0.5/0.5 1.2/1.2 5 6 0
2 1.05/1.05 1.0/1.0 1.09/1.09 26 24 9
2 1.0/1.0t 1.0/1.0 1.09/1.09 3 0 0

“N, - number of individuals scored at all three loci; at two of the three loci; at one locus.
**Total number of individuals scored at this locus shown in parentheses.

tSee text for explanation.

Constructing reasonable selection hypotheses is difficult
because we lack adequate quantitative data on the ecology
and behavior of Neotricula aperta. These snails have a life
span of approximately one year, and one might expect strong
selection for competitive ability during the two month period
of explosive population growth and high juvenile mortality. Dif-
ferences in growth rates have, in fact, been noted between
“‘races’’ and sexes (Davis et a/., 1976) and between localities
(Kitikoon et a/., 1981). Relationships between heterozygosity
and growth have been suggested for many other organisms
including several marine molluscs (See review by Allendorf
and Leary, 1986). Testing this hypothesis, like the previous
one, would require continuous ecobehavioral and demo-
graphic observations in nature and sampling that is very
sensitive to local population structure.

4. Artificial heterozygote deficiencies due to gel scoring er-
rors. We are highly confident of our scoring for the enzymes
on which the reassortment of the original ‘‘racial’’ samples
was based: GAP, PGI, and PEP-3 in the alpha race and gam-
ma race samples and MDH and PEP-3 in the beta race sam-
ple all form clear, distinct bands. On the other hand, some
of the residual heterozygote deficiencies could be explained
by scoring bias for homozygotes: ACP and PEP-4 in Mekong
River taxon 2 and EST-1 and LAP in Mun River taxon 2. ACP
electromorphs were diffuse and never clearly double-banded.
In the case of PEP-4, there were only slight mobility dif-
ferences between five electromorphs, making presumptive
heterozygotes difficult to discern and, therefore, possibly
underestimated. EST-1 banded faintly and diffusely, never be-
ing clearly double-banded. The two LAP electromorphs seen
in the Mun River taxa were very close in mobility and it is
possible that heterozygotes were not recognized. Even with
these minor qualifications scoring bias cannot account for our
observations.

5. Artificial heterozygote deficiencies due to null alleles.
There was no evidence for acommon null allele at any locus
in any of the original ‘‘racial’’ samples: The missing data for
unscorable snails (Tables 4 and 5) did not assume the random
pattern expected for non-lethal null allele homozygotes or
codominant null allele heterozygotes but tended to occur
together on inferior gels. If null homozygosity was lethal at
a locus, this could account for a heterozygote deficiency even
if blank spots, due to other causes, appear on gels. However,
lethal or sublethal null alleles would have to originate by muta-
tion at unrealistically high rates or have unreasonably high

selection coefficients to explain the levels of heterozygote defi-
ciency shown by the loci in our study. It seems highly unlike-
ly that null alleles, lethal or not, could account for the
generalized strong genotypic disequilibria shown by our
data.

6a. Artificial heterozygote deficiencies due to biased sampl-
ing of homozygotes. The possibility that one or more of the
original ‘‘racial’’ populations was genetically subdivided in
some ecobehavioral way at the time of collection was enter-
tained. If, for instance, a snail’s selection of microhabitat is
correlated with its growth and growth rate is in turn associated
with heterozygosity at one or more loci then it could be possi-
ble to simply miss a highly heterozygous portion of the popula-
tion if the sampling protocol is not carefully designed.
However, the general hypothesis (Zouros et a/., 1980) does
not require the degree of genetic differentiation seen in our
data, and for this reason alone we think it unlikely to account
for the pervasive heterozygote deficiencies seen in all three
of our original samples. Again, the very high genetic identity
(D = 0.01) between the original “‘alpha race’ and “‘gamma
race’ samples and the complete lack of intermediacy between
the two Mekong River taxa as we have defined them (Table
4) argues against this less-than-dramatic sort of population
structuring.

6b. Artificial heterozygote deficiencies due to differential
survival of genotypes following collection. Our snails were
maintained in aquaria according to established culture
methods (Kitikoon, 1981a) for about one month following col-
lection and then frozen. Artificial selection due to collecting,
sorting, and live maintenance procedures is always a con-
cern in this type of study. Although we have no quantitative
information on snail mortality during this one month period,
it was not excessive and we doubt artificial selection accounts
for our observations.

7a. Artificial heterozygote deficiencies due to the unsus-
pected presence of cryptic taxa in the allegedly homospecific
samples. The sampling error known as the Wahlund effect
(Wahlund, 1928) is the hypothesis we think best accounts for
the heterozygote deficiencies in the original ‘‘racial’’ samples
and is, of course, the premise on which our sampling reassort-
ment was made. The complete lack of heterozygotes without
concomitant evidence for null alleles or sex-linkage at not one
but several loci strongly support this hypothesis. If the
heterozygote deficiencies in the original samples were real,
we would not expect such a marked reduction in the pattern

STAUB ET AL.: GENETIC VARIATION IN NEOTRICULA APERTA

Table 6. Allele frequencies for 16 loci in four resorted samples of Neotricula aperta, with summary
statistics of genetic variability.”

Mekong River Mun River
locus/allele Taxon 1 Taxon 2 Taxon 1 Taxon 2
Aat (51)** (55) (2) (14)
1.0 1.00 1.00
09 1.00 1.00
Acp (53) (56) (9) (38)
1.1 0.40 0.24 0.11 0.14
1.0 0.42 0.65 0.78 0.75
0.7 0.18 0.11 0.11 0.11
Est-1 (52) (60) (9) (53)
1.0 0.32 0.72
0.93 0.68 0.28 0.17 0.07
0.88 0.78 0.85
0.81 0.05 0.08
Est-2 (43) (59) (8) (51)
1.0 1.00 1.00 1.00 1.00
Est-3 (56) (64) (9)
1.6 0.01
1.4 0.02 0.94 0.99
1.0 1.00 0.98 0.06
Gapt (55) (61) (9) (42)
1.4 1.00 1.00 1.00
1.0 1.00
a-Gpdh (28) (52) (6) (10)
1.0 1.00 1.00 1.00 1.00
Gpit (63) (61) (11) (55)
2.0 0.80
1.0 1.00 0.20 1.00 1.00
Idh (62) (64) (10) (48)
1.0 1.00 1.00 1.00 1.00
Laptt (61) (61) (11) (48)
al 0.6
1.05 0.92
1.0 0.93 0.97 1.00 0.08
08 0.01 0.03
Mdhtt (61) (66) (10) (56)
1.0 1.00 1.00 1.00
05 1.00
Pep-3t/tt (57) (62) (6) (40)
1.2 0.89 1.00
1.11 0.11 0.04
1.09 1.00
1.0 0.96
Pep-4 (59) (64) (9) (54)
1.14 0.08
1.07 0.10 0.48
1.0 0.26 0.29
0.96 1.00 1.00
09 0.35 0.14
0.8 0.29 0.01
Pgd (51) (64) (10) (41)
18 0.47 0.04
16 0.05 0.02
1.0 0.53 0.96 0.85 0.60
0.5 0.10 0.38
Pgm-1 (45) (54) (4) (21)
1.07 0.87 0.95
1.0 0.97 0.15 0.13 0.05
0.9 0.03 0.85

(continued)

100

Table 6. (Continued)

AMER. MALAC. BULL. 7(2) (1990)

Mekong River Mun River

locus/allele Taxon 1 Taxon 2 Taxon 1 Taxon 2
Pgm-2 (50) (55) (11) (31)

-0.7 0.20 0.17

-0.9 0.34 0.10

-1.0 0.46 0.46

1.1 0.27 1.00 1.00
N 529 59.9 8.4 40.2
(S.E.) (2.2) (1.1) (0.7) (3.6)
A 18 2.0 1.5 1.6
P 50.0 625 31.3 37.5
H 0.18 0.15 0.09 0.05

*Samples are described in text; N = mean sample size (and standard error) per locus, A = mean
no. alleles per locus, P = proportion of loci polymorphic, H = mean individual heterozygosity.

**Sample size.

tOne of the loci used to characterize the newly recognized Mekong River taxa.
ttOne of the loci used to characterize the newly recognized Mun River taxa.

following reassortment (Table 2). Instead, we would expect
to see random changes among loci. The same sort of reason-
ing applies to the decrease in the number of loci showing
significant differences in allele frequencies between males
and females. The virtual absence of these sexual differences
in our resorted samples (Table 2) argues that they and not
the original racial samples best represent the natural taxa
present.

7b. Artificial heterozygote deficiencies due to errors in field
identification and the inclusion of several previously recog-
nized taxa in allegedly homospecific samples. We discuss this
hypothesis last as, if correct, it would seriously compromise
our conclusions. Two species found in the Mekong River have
been reported to be similar enough to Neotricula aperta in
size, shell shape, and mantle pigmentation to confuse col-
lectors (Temcharoen, 1971; Davis et a/., 1976). ‘‘Manningiella’’
conica is closely related to or congeneric with Tricula (sensu
lato; Davis, 1979; Kitikoon, 1981b) but so far has only been
found at Khong Island, southern Laos (Davis et al., 1976;
Kitikoon et al., 1981; Kitikoon, 1984), some 200 river km south
of the site where we collected our alpha and gamma race
samples. Moreover, it is most often found by sieving sand,
a substratum not associated with N. aperta (Davis et al., 1976).
It has also been suggested that Pachydrobia bavayi could also
be confused with N. aperta since its young have a shell very
similar to that of ‘‘M’”’ conica (Davis et al., 1976; Upatham et
al., 1983). We think that the possibility of a generic misiden-

Table 7. Matrix of genetic similarity and distance coefficients for four
newly recognized taxa previously referred to Neotricula aperta’.

Sample 1 2 3 4
1 Mekong River taxon 1 --- 0.67 0.56 0.51
2 Mekong River taxon 2 0.34 --- 0.44 0.42
3 Mun River taxon 1 0.54 0.84 -- 0.78
4 Mun River taxon 2 0.68 0.88 0.22 ---

*Below diagonal: Nei’s (1978) unbiased genetic distance (D); above
diagonal: Roger’s (1972) genetic similarity (S).

tification is remote as, in the Mekong River in May,
Pachyarobia sp. are typically 2-3 times the size of N. aperta
and associated with soft-bottom microhabitats (Davis, 1979).

We conclude that the observed heterozygote deficien-
cies were most probably artificial and due to the insensitivity
of our field sampling, and field and laboratory sorting to detect
the previously unrecognized cryptic taxa coexisting at each
site. We accordingly proceed to discuss the significance of
the observed genetic distances between these newly
discovered taxa.

TAXONOMIC INTERPRETATION OF MULTILOCUS
GENETIC DISTANCES

As shown in Figure 1, we have detected two well-
differentiated (D = 0.34) sympatric taxa in the Mekong River.
In the Mun River, we discovered two other well-differentiated
(D = 0.22) sympatric taxa. The newly recognized taxa within
each river are more closely related to one another than to the
taxa in the other river (D = 0.74). These estimates of genetic
differentiation are based on 16 loci and a technique ap-
propriate to establishing evolutionary relationships among
congeneric taxa (Richardson et al/., 1986; Nei, 1987).

At the outset it must be stressed that there is no sim-
ple relationship between a Nei’s genetic distance (D) value
and taxonomic level. Other factors including innate genetic
variability, mating system, effective population size and degree
of population subdivision will all affect the rate of genetic
divergence in a clade. Nevertheless, much can be learned
from the vast literature on genetic differentiation within and
between other well-characterized amphimictic (sexually
reproducing, outcrossing and moderately polymorphic)
species. For example, Thorpe (1983) reviewed over 7000 com-
parisons of conspecific populations of plants and animals and
found that only 2% of the intraspecific D values exceeded 0.10.
In contrast, he found that the average interspecific genetic
distance in 900 congeneric comparisons was about 0.40
(range: 0.03 - >1.0). A survey of 23 genera of amphimictic

STAUB ET AL.: GENETIC VARIATION IN NEOTRICULA APERTA 101

a/y RACE MEKONG R., TAXON 1
a/y RACE MEKONG R., TAXON 2
8 RACE MUN R., TAXON 1
8 RACE MUN R., TAXON 2

1.00 0.50 0

NEI’S GENETIC DISTANCE (D)

Fig. 1. Dendrogram showing relationship of four newly recognized
sibling species previously referred to Neotricula aperta, generated
by UPGMA cluster analysis based on 16 loci.

molluscs revealed they too typically have intraspecific genetic
distances of <0.10 and congeneric interspecific genetic
distances in the range 0.20 - 0.80 (Woodruff et a/., 1988). We
conclude that our estimates of genetic differentiation within
the taxon formerly called Tricula aperta are of such magnitude
that each of the four newly discovered taxa warrant recogni-
tion as separate full species. Our only reservation about this
recommendation arises from the lack of data on intraspecific
variability within each of these sibling species. If, as expected,
intraspecific variation is small (D <0.10), and the gap between
intraspecific and interspecific genetic distances remains
relatively large, then the genetic distance values alone indicate
these taxa are evolving separately as different biological
species.

There is, of course, nothing new about the use of
allozyme electrophoresis to detect sibling species. Bullini
(1983) and Ayala (1983) review the successful use of the
technique in the detection of sibling species in ascarid worms,
plethodontid salamanders, Anopheles mosquitoes and
Drosophila. Other examples involve the Asian schistosomes
transmitted by snails of the genera Tricula, Robertsiella and
Oncomelania (Fletcher et a/., 1980; Woodruff et a/., 1987a;
Merenlender et a/., 1987). Studies of allozyme variation used
in conjunction with traditional methods have been particularly
useful in resolving the evolutionary relationships of tax-
onomically difficult groups of molluscs (Woodruff and Gould,
1980, 1987; Gould and Woodruff, 1986, 1987; Woodruff et a/.,
1987b; Klinhom, 1989; Palmer, Gayron and Woodruff, unpub.
data). Davis (1983, 1984) has used electrophoretic data to
detect sibling species in other molluscs, but did not include
this technique in his early studies of the triculines.

OTHER EVIDENCE THAT NEOTRICULA APERTA
COULD BE A COMPOSITE TAXON

Kitikoon (1982a) described extraordinary variation in
chromosome number and appearance for each on the so-
called races of Neotricula aperta. Haploid chromosome
numbers ranged from 13 to 17 in alpha race males, from 14
to 17 in beta and gamma race males, and from 16 to 17 in
alpha and gamma race females. Diploid chromosome
numbers were 29, 31, and 33 in alpha and gamma race males,
31 and 33 for beta race males, and 32 and 34 in alpha and
gamma race females. Only beta race females showed no
variation with 17 haploid and 34 diploid chromosomes. The

pairing patterns at prophase | were also variable as were other
aspects of the karyotype. This degree of variation within a
single species is almost unknown (White, 1973) and suggests
that Kitikoon’s samples could have been as heterogeneous
as our own. A new analysis based on allozymically sorted
specimens could provide more coherent results.

Kitikoon’s (1982b) electrophoretic study of 5 enzymes
in the three races also revealed considerable interracial varia-
tion, but his results cannot be interpreted genetically as he
pooled tissues of 40-100 snails to prepare his racial samples.

Kitikoon’s work was based primarily on field collections
made in 1972-4 and in 1979. In addition to recognizing the
three so-called races, and noting the occurrence of some
populations that did not conform to this classification, he
distinguished two additional phenotypes of the gamma race
from Sompamit Falls, southern Laos (Kitikoon and Schneider,
1976; Kitikoon et a/., 1981). He subsequently referred to the
latter as (unnamed) separate species (Kitikoon, 1984). Clearly,
both he and his colleagues recognized the complexity of the
taxon called Neotricula aperta.

PARASITOLOGICAL IMPLICATIONS

So far only the gamma race has been unequivocally
shown to transmit Schistosoma mekongi naturally (Kitikoon
et al., 1973), but the alpha and beta races are susceptible to
miracidia infection with subsequent cercarial shedding in the
laboratory (Kitikoon, 1981b; Yuan et a/., 1984). Several authors
have compared rates of snail susceptibility but with inconsis-
tent results (see Kitikoon, 1981b). However, there seems to
be general agreement that, in the laboratory, beta race snails
are highly susceptible, alpha race snails have low suscep-
tibility, and gamma race snails are intermediate with respect
to this trait (Kitikoon, 1981b). As host-parasite compatibility
evolves on a very localized geographic basis in nature (Rollin-
son and Southgate, 1985; Woodruff, 1985), these laboratory
experiments tell us rather little about the potential for the
spread of human schistosomiasis from the transmission site
at Khong Island, Laos. Our findings suggest that the identity
of the intermediate host snail must now be reestablished and
the epidemiological significance of its sibling species
reinvestigated.

CONCLUSIONS

It is now apparent that there are two discrete taxa pre-
sent in the Mekong River that do not coincide genetically with
the so-called alpha and gamma races of Neotricula aperta.
Similarly, in the Mun River we discovered two sibling species
presently confused under the name of the beta race of N. aper-
ta. The genetic distances between these four taxa are large
enough for us to conclude that all have reached the rank of
full species. The lack of evidence for intermediacy in
diagnostic allozyme characters support this. Formal taxonomic
revision must, however, await the confirmation of these pat-
terns by more careful recollection in the field and reexamina-
tion of the anatomy, morphology and karyotypes of the
genetically defined taxa. The type locality of N. aperta is

102 AMER. MALAC. BULL. 7(2) (1990)

Khong Island, Laos, and the holotype conforms to the so-
called alpha race (Davis et a/., 1976). Until snails from this
area can be recollected, it is unlikely that we can resolve the
issues raised by this study of genetic variation in Thai animals.

ACKNOWLEDGMENTS

This study was supported primarily by the United States
Agency for International Development (Program in Science and
Technology Cooperation) and secondarily by grants from the
UNDP/World Bank/WHO Special Programme in Research and Train-
ing in Tropical Diseases, Mahidol University and the University of
California Academic Senate. We thank M. Patricia Carpenter for in-
valuable support in the electrophoresis laboratory and Dr. Viroj
Kitikoon for his comments on the manuscript. Parts of this report were
submitted in partial fulfillment of the requirements for a graduate
degree (Staub, 1988).

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Date of manuscript acceptance: 10 November 1989.

CELLULAR DNA CONTENTS OF THE FRESHWATER SNAIL
GENUS SEMISULCOSPIRA (MESOGASTROPODA: PLEUROCERIDAE)
AND SOME CYTOTAXONOMICAL REMARKS

HIROSHI K. NAKAMURA’
DEPARTMENT OF ZOOLOGY, FACULTY OF SCIENCE
KYOTO UNIVERSITY, KYOTO 606, JAPAN

YOSHIO OJIMA2
DEPARTMENT OF BIOLOGY, KWANSEI! GAKUIN UNIVERSITY
UEGAHARA, NISHINOMIYA 662, JAPAN

ABSTRACT

The cellular DNA contents from eight Japanese Semisulcospira: S. libertina (Gould) and S. reiniana
(Brot) of the S. libertina group, a widely distributed species complex in the Japanese Islands; and S.
decipiens (Westerland), S. habei Davis, S. morii Watanabe, S. multigranosa (Boettger), S. niponica (Smith)
and S. reticulata Kajiyama and Habe of the S. niponica group, the Lake Biwa endemic species com-
plex, were measured by microfluorometry with DAPI staining. Although the species of the S. /ibertina
group (2n=36 and 40) and those of the S. niponica group (2n=14, 24, 26 and 28) had been reported
to have very different chromosome number, the measured DNA values were nearly the same, 3.4 ~
3.7 pg/diploid. Hence it is deduced that karyotypical evolution resulting in different chromosome numbers
between two species groups and within each group could occur without large genomic alternation.

Freshwater prosobranch snails of the genus
Semisulcospira are a diverse and conspicuous element of the
freshwater fauna of the Far East Asia. In Japan they are most
abundant in springs, spring-fed rivers and streams (Kuroda,
1929). The taxa are relegated to two species groups, the
S. libertina group, including widely distributed species in the
Japanese Islands, and the S. niponica group, including many
endemic species of Lake Biwa (Davis, 1969).

There is a wide variation in chromosome numbers
within the genus Semisulcospira. This is remarkable because
constancy of chromosome numbers within large taxa has
been pointed out frequently in various molluscan groups (Pat-
terson, 1969; Nakamura, 1985). In the present study, we
measured the cellular DNA contents of eight species of
Semisulcospira and examined whether the difference of the
chromosome number in the genus could reflect changes in
the genome size. As chromosomal rearrangement and
karyotypical alteration have been thought to be essential in

1Present address and reprint requests: Takasago Research and
Development Center, Mitsubishi Heavy Industries, Ltd., 2-1-1
Shinhama, Arai-Cho, Takasago 676, Japan.

2Present address: Japan Fish Bioscience Institute, 1-17 lwakunicho,
Ashiya 659, Japan.

Semisulcospira speciation (Boss, 1978), data presented here
allow a first assessment of this condition with regard to cellular
DNA contents and chromosomes.

MATERIALS AND METHODS

Specimens of Semisulcospira spp., except S. libertina,
were collected from Lake Biwa in August, 1988 and identified
by N. C. Watanabe. Table 1 shows the localities and dates
of collection. Cells of the embryos in the pallial brood pouch
of two females of each species were prepared and examined
by DNA microfluorometry with DAPI (4, 6-diamidino-2-
phenylindole) staining of Komaru et a/. (1988) with slight
modifications:

1) Crack off the individual adult shells, and entirely dissect
out the pallial brood pouches.

2) Place the individual brood chamber in a separate vial filled
with 0.25% trypsin prepared in calcium and magnesium
free phosphate buffer solution (PBS) and crush the em-
bryonic shells with a glass rod.

3) Allow the trypsin to act on the embryonic body tissue in
the vial over a magnetic stirrer at room temperature for 30
min.

4) Decant the contents into a 15 ml centrifuge tube and spin

American Malacological Bulletin, Vol. 7(2) (1990):105-108

105

106 AMER. MALAC. BULL. 7(2) (1990)

at 100xg for 7 min.

5) Discard supernatant and wash cells once in PBS.

6) Fix in freshly mixed Carnoy’s fixative (3:1 methanol-glacial
acetic acid) by slow addition for 5 min.

7) Centrifuge and change the fixative three times and final-
ly resuspend pellet.

8) Add one drop of cell suspension to a clean washed glass
slide and air dry.

9) Stain the cells left on the slide with the DAPI solution for
1 hr at 49°C.

10) Mount the stained slide with the DAPI solution, cover with
a cover slide, and seal with clear manicure cement. DAPI
solution contained 50ng/ml 4’, 6-diamidino-2-phenylindole
dihydrochloride and 10mM 2-mercaptoethylamine hydro-
chloride in Tris buffer (10mM Tris, 10mM EDTA-2Na,
100mM NaCl, pH 7.4).

Though the absolute cellular DNA concentration for
more than 110 molluscan species was reported by Hinegard-
ner (1974), few of them are available in Japan. Therefore, we
used goldfish (Carassius auratus Temmick and Schlegel) cells
as the standard to estimate absolute DNA amounts of our
snails. The fin epithelium of C. auratus was fixed and prepared
in the same way as mentioned above.

Cell nuclei stained with DAPI were excited by ultra
violet light (365nm) with an Olympus fluorescence microscope
BHS-RFX. The optical conditions were as follows: excitation
filter UGI, dichroic mirror DM400, cut filter L420, objective lens
UVFL 40x.

RESULTS

We were able to obtain very consistent fluorescence
intensity measurements, because the fluorescence from DAPI
stained cells was very intense and mounting the slides with
DAPI solution reduced its decay (Hamada and Fujita, 1983).
The results of DNA estimates of the eight Semisulcospira
species are shown in Table 2. The value relative to the goldfish
standard was converted to an estimate of DNA/nucleus by
multiplying the DNA value in the goldfish nucleus by 4.0
pg/diploid (Hinegardner, 1972). The estimated DNA quantities,
3.4 ~ 3.7 pg/diploid, were not as variable as the chromosome
numbers among the species, 2n=14 ~ 40; and there was lit-
tle interspecific variation.

DISCUSSION

The microfluorometric procedure with DAPI staining is
simple and useful for quantification of DNA content. Recent-
ly, this method has been successfully applied to measure the
ploidy of a variety of molluscs; pearl oyster larvae, Pinctada
fucata martensii (Uchimura, et al., 1987) and scallop, Chlamys
nobilis (Komaru, et al., 1988). It has been demonstrated to
be convenient and sufficiently accurate to be a substitute for
flow cytometry. Using this procedure, we have determined the
nuclear DNA content of eight species of Semisulcospira. The
snails were shown to possess 3.4 ~ 3.7 pg/diploid, which is
within the limits of the DNA values reported previously for the
Mollusca and more specifically the Mesogastropoda. These

Table 1. Collection data of specimens studied here.

Semisulcospira libertina group*
S. libertina (Gould, 1859) Uegahara Waterway (western side
of the Kwansei Gakuin University
campus), Nishinomiya City, Hyogo
Pref., 10 Mar 1988.
Lake Biwa, around Hydrobiological
Station, Kyoto Univ., Otsu City,
Shiga Pref., 5 Aug 1988.

S. reiniana (Brot, 1876)

S. niponica group*
S. decipiens (Westerland, Lake Biwa, from a stretch of the
1883) lake about 1 km off shore (from
Shina to Shimo-sakamoto section),
Shiga Pref., 5 Aug 1988.
Lake Biwa, Konohama Beach,
Moriyama City, Shiga Pref., 5 Aug
1988.
Lake Biwa, Chikubu-jima Island,
Shiga Pref., 3 Aug 1988.
S. multigranosa (Boettger, Lake Biwa, mouth of Kusatsu River,
1866) Kusatsu City, Shiga Pref., 3 Aug
1988.
Lake Biwa, Uchide-hama Beach,
Otsu City, Shiga Pref., 2 Aug 1988.
Lake Biwa, from a stretch of the
lake about 1 km off shore (from
Shimo-sakamoto to Shina section),
Shiga Pref., 5 Aug 1988.

S. habei Davis, 1969

S. morii Watanabe, 1984

S. niponica (Smith, 1876)

S. reticulata Kajiyama and
Habe, 1962

“According to Davis (1969).

values are slightly higher but close to the upper limit (about
3.2 pg/diploid) of the distribution (Hinegardner, 1974).

Davis (1969) subdivided Japanese Semisulcospira in-
to two species groups: the S. /ibertina group and the S.
niponica group. The former includes widely distributed species
in the Japanese Islands and is characterized by larger
chromosome numbers (n=18 or 20), many basal cords and
many embryos per female. The latter consists of several en
demic species of Lake Biwa and is characterized by smaller
chromosome numbers (n=7 to 14), fewer basal cords and
fewer embryos per female. Japanese Semisulcospira show
considerable variety in chromosome numbers (Burch, 1968)
and have therefore attracted our attention, because a general
conservativeness with regard to chromosomal change is evi-
dent in many molluscan groups (Patterson, 1969; Nakamura,
1985, 1986).

In the present investigation we have measured the
cellular DNA amounts to ascertain if the genome can change
among the species as their chromosome number changes.
We have found that the DNA values were very constant; in-
dependent of the divergence in chromosome numbers. Hence
we infer that the difference in chromosome number could oc-
cur without large genomic alteration between the two species
groups and within each group.

Boss (1978) interpreted that the reduction in
chromosome number could have occurred in the karyotype
of the Lake Biwa endemic species as a consequence of
Robertsonian type of chromosomal rearrangement. This

NAKAMURA AND OJIMA: DNA CONTENT OF SEMISULCOSPIRA

Table 2. Cellular DNA contents in Semisulcospira spp. and Carassius auratus.

Species Number of DNA value relative Estimated DNA _ Diploid chromosome
cells examined to the goldfish per cell number*1(2n);
from all indiv. (mean + S.E.) (pg/diploid) arm number (NF)

S. libertina 216 89.0 + 13 3.6 2n=36; NF=72 (66*2)

S. reiniana 198 913 + 09 3.7 2n=20

S. decipiens 174 846 + 09 3.4 2n=24; NF=44

S. habei 251 86.0 + 1.2 3.4 2n=14; NF=28

S. morii 144 866 + 1.0 3.5 2n=32; NF=60*3

S. multigranosa 233 89.9 + 1.1 3.6 2n=28; NF=28

S. niponica 214 876 + 1.0 3.5 2n=24; NF=44

S. reticulata 288 93.6 + 1.1 3.7 2n=24; NF=24

C. auratus 305 100.0 + 08 4.0*4 ——

*Lafter Burch (1968), but NF’s calculated by the present authors.

107

*2-calculated according to the karyotype reported by Kobayashi (1986).
*3-calculated according to the karyotype reported by Watanabe (1984).

*4-after Hinegardner (1972).

phenomenon has been observed in many insect groups (see
White, 1978 for examples) and it is assumed that their DNA
contents remained essentially constant.

Although the constancy of DNA content was confirm-
ed in the present study, something more complex and involv-
ing chromosomal alterations, is probably taking place. This
has also been pointed out by Boss (1978). In the case of
Robertsonian processes, the arm numbers or so-called the
NF’s (‘‘nombre fundamental’ of Matthey, 1945) of the
chromosomes should correspond to the different chromosome
numbers for all species. At the least, there would be much
closer correlation among NF’s than the chromosome
numbers. Table 2 shows that there seems to be no relation-
ship between the chromosome number and the NF in this
group. For situations like this, Matthey (1973) presents some
other possible explanations, especially for lower chromosome
numbers, e.g. tandem fusion of the several smaller chromo-
somes producing one larger chromosome. Much more de-
tailed information on the karyotypes, however, is needed to
determine if such phenomena were applicable to the
karyotypical evolution in the Semisulcospira. The size of each
chromosome complement and other characteristics, such as
banding patterns, to identify homologous chromosomes
among the different species are left for the future studies.

Recently, Kobayashi (1986) reported very different
results on karyotypes than those of Burch (1968), including
chromosome number. One of the Lake Biwa endemic species,
S. nakasekoae, was presented to have not smaller, but slight-
ly larger chromosome number, 2n=38, than S. libertina, a
typical representative of the widely distributed species.
However, as incorrect and inconsistent use of chromosome
terminology was used in the text, figures and tables, the arti-
cle is partially difficult to understand. Watanabe (1984)
reported a new chromosome number, 2n=32, in the genus
from a new Lake Biwa endemic species, S. morii; however,
this result was misquoted by Kobayashi (1986). Although
karyological condition remains very complicated, the present
study leads us to assume that the genome size cannot be
changed largely within the Semisulcospira. This could pro-

vide a basis for investigating the mechanisms of the diver-
sification in this group, and to accumulate more chromosomal
information together with reexamination on the species
previously studied by Burch (1968).

ACKNOWLEDGMENTS

Our acknowledgments are expressed to the staff of the Na-
tional Research Institute of Aquaculture, in particular to A. Komaru
and K. T. Wada for their kindness in providing us with facilities of
microfluorometry and for discussing various aspects of this work. The
specimens from Lake Biwa were collected by M. Nishino, Lake Biwa
Research Institute, and N. C. Watanabe, Kagawa University, and the
computer programs to analyze and compile the data were provided
by T. Hikida, Kyoto University, and M. Takai, Kwansei Gakuin Univer-
sity, to whom our thanks are due for their cooperation. Our thanks
are also expressed to K. Yamamoto, Kwansei Gakuin University, for
excellent technical assistance, and to S. J. Townsley, University of
Hawaii, for critically reading the manuscript. This work was supported
in part by Lake Biwa Research Institute and by a Grant-in-Aid to one
of the authors (HKN), for Scientific Research from the Ministry of
Education, Science and Culture (No. 62790179).

LITERATURE CITED

Boss, K. J. 1978. On the evolution of gastropods in ancient lakes, /n:
Pulmonates. vol. 2A Systematics, Evolution and Ecology, V. Fret-
ter and J. Peake, eds. pp. 385-428. Academic Press, New York.

Burch, J. B. 1968. Cytotaxonomy of Japanese Semisulcospira (Strep-
toneura: Pleuroceridae). Journal de Conchyliologie 107:1-51.

Davis, G. M. 1969. A taxonomic study of some species of Semisul-
cospira in Japan (Mesogastropoda: Pleuroceridae).
Malacologia 7(2/3):211-294.

Hamada, S. and S. Fujita. 1983. DAPI staining improved for quan-
titative cytofluorometry. Histochemistry 79:219-226.

Hinegardner, R. 1972. Cellular DNA content and the evolution of
teleostean fishes. American Naturalist 106(951):621-644.

Hinegardner, R. 1974. Cellular DNA content of the Mollusca. Com-
parative Biochemistry and Physiology 47A:447-460.

Kobayashi, T. 1986. Karyotypes of four species of the genus
Semisulcospira in Japan. Venus 45(2):127-137 (In Japanese with
English abstract).

108 AMER. MALAC. BULL. 7(2) (1990)

Komaru, A., Y. Uchimura, H. leyama, and K. T. Wada. 1988. Detec-
tion of induced triploid scallop, Chlamys nobilis by DNA
microfluorometry with DAPI staining. Aquaculture 69:201-209.

Kuroda, T. 1929. On the species of Japanese Kawanina (Semiscul-
cospira). Venus 1(5):179-193 (In Japanese).

Matthey, R. 1945. ‘evolution de la formule chromosomiale chez les
vertébrés. Experientia 1:50-56 et 78-86.

Matthey, R. 1973. The chromosome formulae of eutherian mammals.
In: Cytotaxonomy and Vertebrate Evolution. A. B. Chiarelli &
E. Chapanna, eds. pp. 531-616. Academic Press, New York.

Nakamura, H. K. 1985. A review of molluscan cytogenetic informa-
tion based on the CISMOCH-Computerized Index System for
the Molluscan Chromosomes. Bivalvia, Polyplacophora and
Cephalopoda. Venus 44:193-225.

Nakamura, H. K. 1986. Chromosome of Archaeogastropoda

(Mollusca: Prosobranchia), with some remarks on their cytotax-
onomy and phylogeny. Publications of Seto Marine Biological
Laboratory 31(3/6):191-267.

Uchimura, Y., A. Komaru, K. T. Wada, H. leyama, H. Yamaki, and
H. Furuta. 1987. Detection of induced triploid larvae of the
Japanese pearl oyster, Pinctada fucata martensii by
microfluorometry with DAPI staining. Fish Genetics and
Breeding Science 12:57-70.

Watanabe, N. C. 1984. Studies on taxonomy and distribution of
freshwater snails, genus Semisulcospira in the three islands
inside Lake Biwa. Japanese Journal of Limnology 45(3):194-203.

White, M. J. D. 1978. Modes of Speciation. Freeman, San Francisco.
455 pp.

Date of manuscript acceptance: 26 July 1989.

USE OF SHELL MORPHOMETRIC DATA TO AID CLASSIFICATION
OF PISIDIUM (BIVALVIA: SPHAERIIDAE)

BRUCE W. KILGOUR
DENIS H. LYNN
GERALD L. MACKIE
DEPARTMENT OF ZOOLOGY, UNIVERSITY OF GUELPH,
GUELPH, ONTARIO, CANADA, N1iG 2W1

ABSTRACT

Univariate and multivariate statistical techniques were applied to 13 shell measurements of clams
from a total of nine populations of four species of Pisidium. Using morphometric data, P compressum
(Prime, 1852) and P subtruncatum (Malm, 1855) can be separated from each other, and from P adamsi
(Stimpson, 1851) and P casertanum (Poli, 1795). P adamsi and P casertanum can be separated using
morphometric data if they are collected from the same location. However, classification of these two
species is difficult when shells from different habitats are compared.

In the Sphaeriidae, there are few discontinuous
variables that can be successfully used to discriminate among
species. As such, identification in the past has been based
on shell shape but using subjective criteria (Herrington, 1962;
Clarke, 1973; Burch, 1975; Mackie et a/., 1980). Since there
tends to be variation in the form of shells among populations
(Holopainen and Kuiper, 1982; Bailey et a/., 1983; Mackie and
Flippance, 1983), and differences between species can be
subtle, there is potential for mis-identification of clams.

Pisidium casertanum (Poli, 1795) is considered by many
authors as the most widespread and common of the
Sphaeriidae. According to Herrington (1962), Burch (1975), and
Holopainen and Kuiper (1982), this species, also, exhibits the
greatest variation in shell form among populations. According-
ly, species that are morphometrically similar to P casertanum
can be difficult to identify correctly. One such species is P
adamsi (Stimpson, 1851). According to Mackie (1989), P
adamsi has a longer dorsal margin that is more gently curved
and has a steeper anterior slope than P casertanum. Using
these characters, these two species are still difficult to
separate. Therefore, it is necessary that more objective means
be used for description of these two species.

Typically, length, height and width are used to objec-
tively describe shell shape in bivalves (e. g. Eager, 1978; Eager
et al., 1984; Mackie, 1989). However, using ratios of these
measurements, shapes of Pisidium casertanum and P. adamsi
are not significantly different from each other (Mackie, 1989).
It is necessary, therefore, to develop new measurements

that will more accurately describe shell shape for sphaeriid
bivalves.

Pisidium compressum (Prime, 1852) and P subtrun-
catum (Malm, 1855) are species that are more easily iden-
tified using shell shape. In this study, 14 morphometric
measurements of five populations of P casertanum, two
populations of P adamsi and one each of P compressum, and
P subtruncatum were collected to determine if shell morpho-
metric data can be used to separate species of Pisidium.

METHODOLOGY

SPECIMEN COLLECTION

Pisidium adamsi, P casertanum, P compressum, and
P subtruncatum were collected from 0.1-1.0 m depths of water
with hand sieves (maximum opening 0.7 mm) from six loca-
tions in Ontario, during May, 1987: Aberfoyle Creek (43°28’N,
80°09’W); Carp River (45°29’N, 76°14’W); Golden Lake
(45934’N, 77921’W); Hanlon Pond (43°33’N, 80°15’W); White
Lake (45934’N, 77°21’W); and Yantha Lake (45°30’N, 77°937’W)
(Table 1). Shell form in the Sphaeriidae is affected by water
hardness (Mackie and Flippance, 1983) and habitat type
(Bailey et a/., 1983). Therefore, P casertanum were collected
from habitats exhibiting a range of hardness and habitat type.
By collecting P casertanum in this manner, it was expected
that the range in possible forms of this species would be ac-
quired, and that the study would discover some character(s)
that could be used to separate all populations of P casertanum

American Malacological Bulletin, Vol. 7(2) (1990):109-115

109

110 AMER. MALAC. BULL. 7(2) (1990)

Table 1. Location of populations of Pisidium in Ontario, Canada.

Population Location Code Cal N2
P adamsi
Carp River Fitzroy Twp., Carleton Co. a 230 27
White Lake Bagot and McNab Twps., b 95 18

Renfrew Co.
P casertanum

Aberfoyle Puslinch Twp., Wellington Cc 185 22
Creek Co.

Carp River Fitzroy Twp., Carleton Co. d 230 33

Golden Lake North and South Algona e 34 19

Twps., Renfrew Co.
Guelph Twp., Wellington Co. f 292 25
Sherwood Twp., Renfrew Co. g 44 14

Hanlon Pond
Yantha Lake
P compressum

Carp River Fitzroy Twp., Carleton Co. h 230 30
P subtruncatum
Carp River Fitzroy Twp., Carleton Co. i 230 23

1 Calcium hardness (mg CaCQ3I-’).
2 Number of clams measured.

from P adamsi.

It is more difficult to identify species of clams with
smaller individuals, so all available size classes of clams were
represented in the study, when possible (Table 2).

MORPHOMETRIC MEASUREMENTS

All measurements of clams were determined using a
binocular microscope equipped with a Bioquant Hipad®
digitizer. For measurement of features of the lateral aspect
of the shell (Fig. 1A), clams were placed on the right valve
in sand such that the dorsal margin was parallel to the first
cross-hair (CH1) of the microscope ocular, which went through
the most posterior projection of the shell. The second cross-
hair (CH2) went through the middle of the umbone. This orien-
tation facilitated measurement of the linear characters A-E and
the areas of quadrants 1-4 (Q1-Q4). For the purposes of this
study, the point at which the two cross-hairs meet will be
termed the centre of the clam. For measurements of the cross-
sectional aspect of the shell (Fig. 1B) clams were re-oriented
in sand with the anterior end projected upwards, such that
the first cross-hair (CH1) went through the maximum width
and the second cross-hair (CH2) bisected the two valves. This
orientation facilitated measurement of the maximum width and
the areas of quadrants 5 and 6 (Q5 and Q6).

The linear measures A-E (Fig. 1A) were taken from the
centre of the cross-hairs to the perimeter of the shell along
the associated cross-hair. Measurement A provides an
estimate of the position of the umbone relative to the posterior
margin of the clam. Measurement B estimates the vertical
position of the posterior projection relative to the umbone.
Measurement C is the maximum distance from the centre of
the clam to the shell margin. Measurement D provides an
estimate of how ‘“‘undercut”’ (Herrington, 1962) the posterior-
ventral corner of the shell is; clams that have shorter distances
are more undercut. Measurement E provides an estimate of
how tapered the anterior-dorsal margin is; clams with shorter
distances are more tapered.

DORSAL

POSTERIOR ANTERIOR

Fig. 1. (A) Lateral view of right valve showing measurements made
on each specimen of Pisidium. (B) Cross-sectional aspect of the shell,
viewed from anterior end, showing measurements made on each
specimen. Letters denote measurements: A, distance from umbone
to posterior margin; B, distance from centre of the clam to the dorsal
margin; C, maximum distance from centre of the clam to the shell
perimeter; D, distance from centre of the clam to the ventral margin;
E, distance from centre of the clam to the anterior margin; Q1-Q6,
quadrants 1 to 6 for which area measurements were made; CH1 and
CH2, cross-hairs one and two.

Table 2. Summary of shell length measurements (mm) of nine popula-
tions of Pisidium from Ontario, Canada.

Population’ Minimum Maximum Mean Std Error
a 1.597 4.901 3.042 0.139

b 1.999 4643 3.273 0.166

Cc 1.968 5.474 3.640 0.231

d 1.898 4.329 3.387 0.111

e 1.774 3.327 2.563 0.099

f 1.658 3.276 2.538 0.083

g 1.766 5.036 3.172 0.207

h 1.807 3.965 2.854 0.102

i 2.133 4.248 3.337 0.132

1See Table 1 for species, location of each population, and sample size.

Estimates of the area of each of the six quadrants were
also obtained. These estimates indicate how tapered or round-
ed a particular quadrant is; clams with a smaller quadrant
are more tapered in that quadrant while those with a larger
quadrant are more rounded.

KILGOUR ET AL.: SHELL MORPHOMETRICS OF PIS/DIUM 111

STATISTICAL PROCEDURES

Prior to statistical analyses, all data were logio
transformed to improve normality and linearity of relationships.
For morphometrics to be useful for classification they must
compensate for allometric relationships and be able to
separate all size classes of one species from all size classes
of other species. For these reasons, each logo transformed
variable was regressed on logo transformed length. These
among-groups residuals (Reist, 1985, 1986) of individual
measurements were used to describe shape or morphometric
characters of individual clams, and to remove the effects of
size. These residuals were then used in all subsequent
statistical procedures.

ANOVA and Duncan’s multiple range test were per-
formed on the residuals of each measurement to determine
if any single characters could be useful for classifying in-
dividuals. MANOVA established that signficant differences in
population centroids existed (p < 0.0001). Since these existed,
it justified the use of multiple comparisons techniques (i.e.
canonical variates, discriminant functions, and Mahalanobis’
distances) to elucidate further morphometric relationships
among populations of clams.

Canonical variates analysis (CVA) was used to de-
scribed axes of variation that provided maximum discrimina-
tion among populations of clams (Blackith and Reyment,
1971). Plots of population centroids for each canonical variate
and associated 95% confidence ellipses (Altman, 1978) were
used to visualize morphometric differences among popula-
tions. Canonical variates describing axes of shell variability
that accounted for greater than 10% of the variation in the
data set and with eigenvalues greater than 0.5 were con-
sidered meaningful. Variables with large standardized coef-
ficients (i.e. value of coefficient no smaller than one half the
value of the largest coefficient in that variate) and with total
canonical structure coefficients greater than 0.5 were con-
sidered to be important in determining shell shape in that
variate.

Discriminant functions analysis assesses the validity
of the group classifications through a jack-knifing technique
(Blackith and Reyment, 1971). This technique was used to
calcuate discriminant functions for each population of clams.
Each individual clam was then re-classified according to these
discriminant functions. Clams that were re-classified into their
original population were considered correctly re-classified.
Those clams re-classified into a different population were con-
sidered mis-classified. This analysis effectively assessed the
likelihood of making correct classifications using mor-
phometric data. The final procedure was a calculation of
Mahalanobis’ distances that provided a measure of the mor-
phological distance between groups based on the characters
measured (Blackith and Reyment, 1971).

RESULTS

Univariate tests indicated that Pisidium compressum
and P. subtruncatum were significantly different from all other
populations with respect to five of the thirteen morphometric
characters (Table 3). P compressum from Carp River (popula-

tion h) were wider, higher, had a lower posterior projection (B)
and had larger Q1, Q5, and Q6 than any other population.
P. subtruncatum from Carp River (population i) had a higher
posterior projection (B), longer distance from the centre of the
clam to shell margin (C), and had a larger Q3 than any other
population.

There were no measurements that could separate all
populations of Pisidium casertanum from both populations of
P adamsi. However, in Carp River, P adamsi (population a)
was higher, had a longer distance from the centre of the clam
to the shell margin (C), and a smaller Q5 than P casertanum
(population d).

There were three meaningful axes of variation
described by CVA for shells of Pisidium among the nine
populations (Table 4). The first canonical variate (CV)
described an axis of variation from shells that are low and
narrow to shells that are high and wide. The second CV
described an axis of variation from shells that have a round-
ed quadrant 1 (large Q1) and a short anterior-ventral margin
(short measurement C) to shells that have a tapered quadrant
1 (small Q1) and a long anterior-ventral margin (long measure-
ment C). The third CV described an axis of variation from
shells with a long posterior end (long measurement A) to shells
with a short posterior end (short rneasurement A).

Plots of canonical variate centroids and associated
95% confidence ellipses showed that Pisidium compressum
(population h) and P subtruncatum (population i) were
morphometrically different from each other and from all other
populations (Figs. 2-4). P adamsi in Carp River (population
a) was significantly different from P casertanum in Carp River
(population d) (Fig. 3). Separation of both populations of P
adamsi (populations a and b) from all populations of P caser-
tanum (populations b-g) only occurred when canonical
variates 1 and 3, or 2 and 3 were considered together (Figs.
3, 4). P casertanum from Golden Lake (population e) were
separated from all other populations of P casertanum by
canonical variate three (Figs. 3, 4).

From the discriminant functions analysis, 53% of the
re-classified clams had a length that exceeded the mean
length of the population from which they were originally
classified. Over 95% of Pisidium compressum (population h)
and P subtruncatum (population i) were re-classified correct-
ly (Table 5). The majority of P casertanum were re-classified
into their original population or were re-classified into other
populations of P casertanum. Only 5.6% of P adamsi from
White Lake (population b) were re-classified with the Hanlon
Pond population (f) of P casertanum. |n contrast, 18.5% of
P. adamsi from Carp River (population a) were re-classified
into populations c and e of P casertanum (Table 5).

Mahalanobis’ distances (Table 6) indicated that
Pisidium compressum (population h) and P. subtruncatum
(population i) were significantly different, morphometrically,
from each other and from all other populations of clams. Also,
all populations of P casertanum were significantly different
from both populations of P adamsi, except P casertanum from
Yantha Lake (population g), which was not significantly dif-
ferent from P adamsi from Carp River (population a).

112

AMER. MALAC. BULL. 7(2) (1990)

Table 3. Results of the Duncan’s test on shell measurements of nine populations of Pisidium in Ontario, Canada. Population means
sharing the same letter superscript in the same row are not signifying different (p <0.05). Analysis is based on residuals of 13 mor-
phometric measurements after variation due to shell length was removed. Refer to text for explanation of abbreviations.

Variable Population!
a b Cc d e f g h i
vw WX xy y Vv xy Wwxy u xy
Height 0.0006 -0.005 -0.012 -0.013 0.006 -0.010 -0.006 0.038 -0.008
Wxy yz wxyz Ww Vv XyZ WX u z
Width -0.010 -0.026 -0.016 -0.003 0.020 -0.021 -0.007 0.069 -0.028
vw U w Vv x Vv Vv vw Xx
A 0.003 0.019 -0.004 0.004 -0.014 0.007 0.004 -0.001 -0.017
WX x wx Wx Vv Ww w u y
B -0.009 -0.022 -0.007 -0.010 0.025 0.001 0.007 0.049 -0.041
vw y WX Ww Vv y xy Vv u
Cc 0.003 -0.013 -0.002 -0.005 0.005 -0.011 -0.008 0.008 0.019
U U U U U U u u u
D 0.001 -0.006 -0.005 0.002 -0.010 -0.009 -0.001 0.012 0.007
uvw vw uv v U uv uv Ww uv
E -0.006 -0.014 0.006 0.019 0.013 0.004 0.005 -0.026 0.001
w Vv w vw vw Vv Vv U x
Q1 -0.012 0.008 -0.012 -0.001 0.004 0.011 0.010 0.032 -0.044
Vv U vw vw Ww vw vw 7 vw
Q2 0.011 0.041 -0.016 -0.005 -0.024 -0.003 -0.005 0.013 -0.010
Ww WX WX Wx w Xx WX Vv u
Q3 -0.001 -0.009 -0.021 -0.017 0.001 -0.030 -0.023 0.033 0.056
uv uv uv vw u uv uv uv w
Q4 0.011 0.001 0.028 -0.030 0.046 0.029 0.031 -0.005 -0.078
xy y vw w uv WX Ww U y
Q5 -0.038 -0.080 0.018 0.015 0.061 -0.015 0.013 0.077 -0.073
w Ww w w Vv w Ww u w
Q6 -0.014 -0.020 -0.029 -0.021 0.025 -0.031 -0.018 0.113 -0.034

1See Table 1 for species, location of each population, and sample size.

Table 4. Standardized and total structure coefficients for shell morphometric data from nine populations of Pisidium in On-
tario, Canada. Analysis is based on residuals of 13 shell morphometric measurements after variation due to shell length was
removed. The eigenvalue and the proportion of variance accounted for by a particular variate are also presented. Refer to
text for explanation of abbreviations.

Variable Canonical Variate
Standardized Coefficients Total Structure Coefficients

CV1 Cv2 CV3 Cv1 Cv2 CV3
height 1.203 -1.680 -1.134 0.932 -0.028 0.096
width 0.712 -0.121 0.437 0.854 0.158 0.391
A 0.334 0.190 -0.869 0.007 0.452 -0.760
B -0.269 1.727 0.714 0.654 0.440 0.331
C 0.147 -1.053 0.056 0.261 -0.733 0.453
D 0.154 -0.126 -0.171 0.129 -0.122 0.011
E -0.345 0.209 0.223 -0.338 0.097 0.267
Q1 0.150 -0.870 -0.120 0.438 0.521 -0.093
Q2 -0.050 0.340 0.141 0.133 -0.006 -0.460
Q3 0.014 -0.203 0.102 0.324 -0.640 0.129
Q4 0.086 0.252 -0.203 0.022 0.350 0.024
Q5 -0.190 0.327 0.464 0.389 0.362 0.568
Q6 0.290 0.278 0.154 0.846 0.060 0.216
eigenvalue 3.016 1.502 0.745

% variance 0.534 0.265 0.132

KILGOUR ET AL.: SHELL MORPHOMETRICS OF PIS/DIUM 113

DISCUSSION

SEPARATING SPECIES OF P/S/DIUM

Size of clams used does not appear to have affected
the results of this study. Greater than 50% of the misidenti-
fied clams, from the jack-knifed classification, were larger than
the average-sized clams. This suggests that small clams can
be used in a study of this type without resulting in appreciable
size-related bias.

The morphometric measurements that have been
described in this study appear to be useful for separating
species of Pisidium. Both univariate and multivariate statistical
techniques were successful at separating P compressum and
P. subtruncatum from each other and from P adamsi and P
casertanum. In the future, these new measurements, coupled
with adjustments for shell length (i.e. generation of residual
values after length variation is removed), could be useful for
objectively describing shell shape to identify other species
of Pisidium.

In addition to being useful for classifying species with
unique forms, these morphometric data appear to be useful
for separating populations of the same species. For exam-
ple, plots of canonical variate ellipses suggests that P caser-
tanum from Golden Lake (population e) have significantly dif-
ferent forms when compared to other populations of P
casertanum.

CANONICAL VARIATE 2
c/aQi

-2 4 0 1 2 3 4 5
CANONICAL VARIATE 1
height, width —~

Fig. 2. Plot of centroids for canonical variates 1 and 2, with 95% con-
fidence ellipses for 9 populations of Pisidium in Ontario, Canada. Let-
ters denote populations: a and b, P adamsi; c-g, P casertanum; h,
P compressum,; i, P subtruncatum. Arrows indicate direction in which
the variables increase. Refer to text for explanations of abbreviations
of morphometric variables.

CANONICAL VARIATE 3

-2 -1 fc) 1 2 3 4 5
CANONICAL VARIATE 1
height, width _—~
Fig. 3. Plot of centroids for canonical variates 1 and 3, with 95% con-
fidence ellipses for 9 populations of Pisidium in Ontario, Canada. Let-
ters denote populations: a and b, P adamsi; c-g, P casertanum; h,
P compressum, i, P subtruncatum. Arrows indicate direction in which

the variables increase. Refer to text for explanations of abbreviations
of morphometric variables.

SEPARATING PIS/IDIUM ADAMSI AND
P CASERTANUM

The analysis showed that Pisidium adamsi and P
casertanum have unique forms but it is difficult to make con-
fident classifications for all clams using morphometric data.
Also, PR adamsi and P casertanum are more easily
distinguishable when compared within a habitat than among
habitats. For example, univariate tests showed that P adamsi
and P casertanum from Carp River (populations a and d
respectively) can be distinguished from each other using mor-
phometric data. However, there were no single measurements
that could separate both populations of P adamsi from all
populations of P casertanum. Plots of canonical variate
ellipses showed that, in general, P adamsi have a longer
posterior end, a more tapered Q1 and are higher. However,
discriminant functions analysis showed that although most
clams can be correctly classified (e.g. P casertanum from
Aberfoyle Creek and Golden Lake were correctly re-classified
as P casertanum 100% of the time), there are some clams
(e.g. P adamsi from White Lake were incorrectly re-classified
as P casertanum 18.5% of the time) that are difficult to classify
using morphometric data (Table 5). As well, Mahalanobis’
distances showed that the Yantha Lake population of P caser-
tanum (population g) is not morphometrically distinct from
either P casertanum or P adamsi populations from Carp River
(populations d and a respectively), suggesting that there are
forms that could be mistaken for either of P adamsi or P caser-
tanum using morphometric data. It is not reliable, therefore,

114 AMER. MALAC. BULL. 7(2) (1990)

CANONICAL VARIATE 3
A

-4 -3 =2 -1 ie) 1 2
CANONICAL VARIATE 2
—c/a—

Fig. 4. Plot of centroids for canonical variates 2 and 3, with 95% con-
fidence ellipses for 9 populations of Pisidium in Ontario, Canada. Let-
ters denote populations: a and b, P adamsi; c-g, P casertanum; h,
P compressum,; i, P subtruncatum. Arrows indicate direction in which
the variables increase. Refer to text for explanations of abbreviations
of morphometric variables.

to use shell morphometric data to separate P adamsi and P
casertanum.

Although these data show that the morphometric
measurements described here are useful for making a less
subjective classification of some species of Pisidium, some
species such as P adamsi and P casertanum require addi-
tional information. In other molluscs, classification has been

Table 5. Jack-knifed classification, using shell measurements, of nine
populations of Pisidium in Ontario, Canada. Analysis is based on
residuals of 13 shell morphometric measurements after variation due
to shell length was removed first. Values in parentheses refer to the
proportion of mis-classified clams made into each of the indicated
populations.

Species Popula- % correct % clams mis-classified
tion! in other population
P adamsi a 59.3 b (18.5), c (7.4), e (11.1),
i (3.7)
b 778 a (16.7), f (5.6)
P casertanum Cc 59.1 d (18.2), e (9.1), f (9.1),
g (4.6)
d 445 a (6.1), c (18.2), 3 (3.0),
f (12.1), g (9.1), i (6.2)
e 84.2 c (10.5), g (5.3)
f 48.0 b (8.0), c (4.0), d (4.0),
e (4.0), g (32.0)
g 28.6 b (7.1), c (7.1), d (14.3),
e (14.3), f (28.6)
P compressum h 96.7 g (3.3)
P. subtruncatum i 95.7 a (4.3)

1See Table 1 for location of each population and sample size.

Table 6. Mahalanobis’ distances, using shell measurements, between populations of Pisidium in Ontario, Canada. Analysis is based
on residuals of 13 shell morphometric measurements after variation due to shell length was removed.

Population! b Cc d e f g h i
* +s we ae = ne ae ae

a 1.94 2.24 2.18 2.67 2.18 1.99 4.16 3.24
b 3.05 2.81 3.68 2.28 2.48 4.70 4.17
ns ae ns ns a oe

Cc 1.50 2.05 1.35 1.20 5.40 3.78
** * ns a* ak

d 2.25 1.60 1.37 5.10 3.73
e 2.45 1.98 4.21 4.20
ns ae we

f 0.72 5.24 4.38
g 4.75 4.27
h 5.40

1See Table 1 for species, locations of each population, and sample size.

* = Populations are significantly different at P < 0.05.
** = Populations are significantly different at P < 0.01.
ns = Populations are not significantly different.

KILGOUR ET AL.: SHELL MORPHOMETRICS OF PIS/DIUM 115

improved through study of soft part anatomies (e.g. Davis and
da Silva, 1984; Dillon, 1984; Kat, 1983). It would be beneficial
if more detailed studies of the morphologies of P adamsi and
P casertanum could be performed so that more reliable means
of classification be determined. Also, similar shell mor-
phometric studies on all species of Sphaeriidae are needed
before the use of shell morphometric measurements can be
fully assessed.

ACKNOWLEDGMENTS

Comments from Dr. Doug Noltie on an earlier presentation of
this work are greatly appreciated. Dr. Robert Bailey was helpful with
the SAS programs that were used to generate the confidence ellipses.
Comments from two anonymous referees improved the manuscript.
This project was funded by a NSERC Canada operating grant (no.
A-9882) and Employment and Immigration Canada grants to one of
us (G.L.M.).

LITERATURE CITED

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Academic Press, New York. 412 pp.

Burch, J. B. 1975. Freshwater Sphaeriacean Clams (Mollusca:
Pelecypoda) of North America. Revised Edition. Malacological
Publications (Hamburg, Michigan). 96 pp.

Clarke, A. H. 1973. The freshwater Mollusca of the Canadian Interior
Basin. Malacologia 13:1-509.

Davis, G. M. and M. C. P. da Silva. 1984. Potamolithus: Morphology,
convergence, and relationships among hydrobioid snails.
Malacologia 25:73-108.

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and multivariate analysis used to reduce the non-genetic com-
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of some living and fossil bivalve molluscs. Biological Review
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between shape, weight and thickness of shell in four popula-
tions of Venerupis rhomboides (Pennant). Journal of Molluscan
Studies 50:19-38.

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(Mollusca: Pelecypoda). Miscellaneous Publications Museum
of Zoology University of Michigan No. 118. 74 pp.

Holopainen, I. J. and J. G. J. Kuiper. 1982. Notes on the morphometry
and anatomy of some Pisidium and Sphaerium species
(Bivalvia, Sphaeriidae). Annales Zoologica Fennici 19:93-107.

Kat, P. W. 1983. Genetic and morphological divergence among
nominal species of North American Anodonta (Bivalvia:
Unionidae). Malacologia 23:361-374.

Mackie, G. L. 1989. Biology of Freshwater Corbiculacean Clams of
North America. Canadian Governmental Publication Centre,
Hull, Quebec. In press.

Mackie, G. L., D. S. Zdeba and T. W. White. 1980. A Guide to
Freshwater Mollusks of the Laurentian Great Lakes with Special
Reference to the Pisidium. United States Environmental Pro-
tection Agency EPA-600/3-80-068. 144 pp.

Mackie, G. L. and L. A. Flippance. 1983. Relationships between buf-
fering capacity of water and the size and calcium content of
freshwater molluscs. Freshwater Invertebrate Biology 2:48-55.

Reist, J. D. 1985. An empirical evaluation of several univariate methods
that adjust for size variation in morphometric data. Canadian
Journal of Zoology 63:1429-1439.

Reist, J. D. 1986. An empirical evaluation of coefficients used in
residual and allometric adjustment of size covariation. Canaa-
ian Journal of Zoology 64:1363-1368.

Date of manuscript acceptance: 12 June 1989.

POLYMORPHISM FOR SHELL COLOR IN THE ATLANTIC BAY SCALLOP
ARGOPECTEN IRRADIANS IRRADIANS (LAMARCK) (MOLLUSCA:BIVALVIA)

ON MARTHA’S VINEYARD ISLAND

J. A. ELEK' AND S. L. ADAMKEWICZ
GEORGE MASON UNIVERSITY
FAIRFAX, VIRGINIA 22030 U.S.A.

ABSTRACT

Populations of the Bay Scallop, Argopecten irradians irradians (Lamarck, 1819) on the island
of Martha’s Vineyard, Massachusetts, are highly polymorphic for shell colors and patterns. Juvenile
and adult scallops were sampled from natural populations in two ponds on the island and a system
was devised for classifying the wide range of variation found in their shell colors and patterns. This
classification, which we propose as a guide for future genetic analysis, recognizes three background
colors (white, yellow, and orange) and six overlying colors that contribute to a variety of patterns. Fre-
quencies for some of these shell characters differ significantly between ponds and sometimes between
age classes within a pond. For background color, approximately 94% of the entire sample were white,
5% were yellow, and 1% were orange. This polymorphism, which is known to be genetic with rare
yellow and orange alleles dominant to white, appears to be persistent and can be maintained by fre-
quency dependent selection through predation by teleost fish and shore birds. The entire suite of poly-
morphic shell characters could be an instance of hyperpolymorphism maintained by reflexive

selection.

Scallops are renowned for their brilliant shell colors,
but few studies have investigated this characteristic. Abbott
(1954) described the occurrence of several different colors in
shells of the bay scallop Argopecten irradians irradians
(Lamarck, 1819). Clarke (1965) also noted that various shell
colors existed in this species. Using museum collections, he
estimated the frequency of shells with white lower valves in
populations along the Atlantic coast of North America and
showed that significant differences occur for this pattern
variant. Clarke (op. cit.) also remarked that only white shells
occurred in frequencies high enough to record. Kraeuter et
al. (1984) first produced evidence, by using mass spawnings,
that shell colors in A. i. irradians were independent of the en-
vironment, and Adamkewicz and Castagna (1988) have shown
that background color in these scallops is inherited as a single
gene with orange and yellow alleles dominant to white. Their
work has demonstrated the need of a system for classifying
the shell colors to facilitate planning and interpretation of
further breeding experiments and to permit systematic obser-
vation of natural populations.

Many marine mollusks are highly polymorphic and

1Present address: CSIRO, Division of Entomology, PO. Box 1700,
Canberra City, ACT, Australia, 2601.

researchers have investigated whether their shell colors were
determined by genetic or environmental factors. Environmen-
tal control of shell color has been proposed for the gastropods
Turbo cornutus Lightfoot (Ino, 1949), Haliotis rufescens Swain-
son (Leighton, 1961), and Austrocochlea constricta Lamarck
(Creese and Underwood, 1976) as well as for the clam Donax
denticulatis Linné (Wade, 1968). If most such polymorphisms
had an environmental cause, then they would not be suitable
for studies of natural selection. However, several investigators
have now succeeded in demonstrating a genetic basis for
variations in shell color in marine mollusks, and some studies
have linked these variations to differential survival.

Geisel (1970) and Reimchen (1979) have described
situations where variations in shell color of the limpet Acmea
digitalis Rathke and the snail Littorina mariae Philippi re-
spectively, which had appeared to be related to environmen-
tal factors, were actually polymorphisms maintained by selec-
tive predation. A genetic basis for shell color has now been
demonstrated in the mussel, Mytilus edulis Linné (Innes and
Haley, 1977) and the color variants have been shown to dif-
fer in growth rate (Newkirk, 1980) as well as in resistance to
high temperatures (Mitton, 1977). Cole (1975) has shown that,
although it can change during growth under the influence of
environmental factors such as diet, shell color in the

American Malacological Bulletin, Vol. 7(2) (1990):117-126

117

118 AMER. MALAC. BULL. 7(2) (1990)

gastropod Urosalpinx cinerea Say is inherited directly Palmer
(1985) has demonstrated that a single gene determines shell
color in the snail Thais (=Nucella) emarginata Deshayes while
Etter (1988) has shown a relationship between shell color and
survival at high temperatures in the snail Nucella /apillus Linné.

The present study was designed to ascertain the ex-
tent of polymorphism in the scallop Argopecten irradians ir-
radians and the suitability of the variation for ecological and
evolutionary studies. We propose a system for classifying the
very extensive variation observed into a manageable number
of discrete categories or phenotypes. This system has been
used to estimate proportions of these phenotypes in natural
populations from Martha’s Vineyard Island, Massachusetts,
and to compare phenotypic frequencies both between popula-
tions from two different ponds and between adults and
juveniles within a single pond. Although the forces acting on
this polymorphism are not yet Known, the system appears
promising for future study.

MATERIALS AND METHODS

STUDY AREA

The island of Martha’s Vineyard (Fig. 1) is situated
13 km south of Cape Cod, Massachusetts, at 40°N, 70°W. It
is approximately 32 km long and 14 km wide with numerous
large estuarine ponds which support extensive shellfish beds.
Samples were taken from two of these ponds, Lagoon and
Nashaquitsa. Lagoon Pond is on the north shore of the island
and covers about 236 hct with a narrow opening into Vineyard
Haven Harbour. Its salinity ranges from 22 to 34 ppt during
the year, and the seaward portion has numerous scallop beds.
Nashaquitsa Pond covers about 40 hct and, although it is on

me Martha's
Vineyard

ATLANTIC
OCEAN

&

NANTUCKET SOUND

Lagoon Pond

VINEYARD SOUND
Edgartown +

J wee

Wisnitan Pond o

—

ATLANTIC OCEAN

Martha's Vineyard

Fig. 1. Map of Martha’s Vineyard showing the locations of the two
ponds and the position of the island reiative to the eastern coast of
the United States.

the southwest side of the island, it connects through
Menemsha Pond to Vineyard Sound on the northwestern
shore. Salinity ranges between 26 and 35 ppt. Despite its poor
water circulation, the scallop beds are very productive.
Because it was not feasible to sample the ponds ran-
domly, four sampling sites were chosen from each pond to
represent, so far as possible, the range of variation in physical
parameters such as depth, salinity and distance from the
outlet. Sites in both ponds ranged from 0.5 to 2 m in depth,
salinity was from 32 to 33 ppt, and water temperature was
13°C in Nashaquitsa Pond and 13° to 15°C in Lagoon Pond
at time of sampling in May 1984. The substrata at three of
the four sites in each pond consisted of mud with eelgrass
and algae (Gracillaria sp. and Codium spp.). One site in Nasha-
quitsa Pond was muddy with many dead shells, while one site
in Lagoon Pond was sandy and also had many dead shells.

SAMPLING AND SHELL PREPARATION

The scallops were collected by dredging four sites in
each pond. This produced a sample of 302 adults from
Lagoon Pond and 310 adults from Nashaquitsa Pond. The soft
parts were removed and the shells were soaked in strong
detergent, then scrubbed to remove light fouling. A few shells
were soaked in diluted bleach for about 15 min to remove
more persistent fouling, but this treatment did not affect the
underlying shell material or color. Finally, the shells were
sprayed with clear acrylic to restore their ‘‘wet’’ appearance.
During the summer of the same year, juvenile scallops were
sampled by means of spat collectors, synthetic mesh bags
which provided setting sites for the scallop larvae. Bags were
suspended at one site in each pond for the three summer
months of June, July and August. When removed at the end
of August, the bags from Lagoon Pond contained 231 juvenile
scallops and the bags from Nashaquitsa Pond yielded 371.
Because no fouling organisms had settled on them, the young
scallops required no cleaning to reveal shell colors.

Each shell was measured with calipers to the nearest
0.1 mm while the two valves were held closed. Three dimen-
sions were measured as follows: height, greatest distance
from hinge to growing edge, usually taken along the middle
rib; width, greatest distance across ribs, at right angles to the
middle rib; depth, greatest distance through the shell from
top to bottom valve. For the shells of adults, the measurements
of height and depth were combined to produce an index ex-
pressing the degree of curvature of the valves as follows:
relative depth = depth/height.

The classification system for shell color was devised
from a preliminary sample of 250 adult shells taken from a
third pond on Martha’s Vineyard in May of 1983. Reference
shells were selected for each of the colors and patterns to
standardize classification of shell phenotypes, and these
reference shells are held at George Mason University with
one of the authors (S.L.A.). [A selection of these shells can
be seen in color photographs in Elek (1985) and Adamkewicz
and Castagna (1988)].

Because the top and bottom valves differ markedly in the
distribution of pigments, phenotypes were recorded as either
present or absent separately for the top and bottom valves

ELEK AND ADAMKEWICZ: SHELL COLOR IN ARGOPECTEN 119

of each scallop collected. Intensity of pigmentation was not
recorded at all due to the difficulty of making objective
measurements, and extent of coverage on the shell was
recorded only qualitatively. The patterns and colors on each
adult shell were assessed separately in three zones which
corresponded to the length of early juveniles that attach to
eel grass and other raised locations (about 5 mm), of late
juveniles that release from the eel grass to settle on the bot-
tom (about 20 mm), and of fully adult scallops that have stayed
on the pond bottom resting on their lower (right) valves for
nearly a year (over 40 mm).

All statistical analyses were performed using the
Statistical Package for Social Science (SPSS) Version 9 (Nie
et al., 1975). Contingency tests were used to search for signifi-
cant variation among phenotypic frequencies and analysis
of variance was used to test for significant differences in shell
size among the various phenotypes. A result was considered
significant when the probability of its occurrence was less than
0.05 and was considered highly significant when the probabili-
ty was less than 0.01.

RESULTS

CLASSIFICATION OF SHELL COLOR AND PATTERN

The overall appearance of the shell depends on three
factors which are summarized with their individual variants
in Table 1. These factors, the background color of the shell,
the pattern applied over this ground color, and the color(s)
used to produce the patterns, are definitely not independent
of one another in their occurrence, and only for background
color is the genetic basis known. The analysis of the patterns
and their colors is further complicated because these two fac-
tors have a strong developmental component. Both the in-
teractions and the developmental sequence will be described
after the basic variants of the factors and their frequencies
have been presented.

Background color is defined as the color which suf-
fuses both valves of the shell and which is produced uniformly
throughout the life of the animal (i.e. intensity does not vary).
Three background colors were identified which were mutual-
ly exclusive in their occurrence: 1) white, probably an absence
of pigment; 2) yellow, ranging in intensity from cream to
golden; 3) orange, also varying in intensity. Scallops always
had one of the three alternative background shell colors, the
same on both valves.

Pattern colors are pigments overlying the background
color and not covering the entire shell. Unlike background
color, pattern colors are not alternatives and can be different
on top and bottom valves. Six different colors were identified
which could occur either separately or in any combination with
one another and could overlay any of the three background
colors. These pattern colors were further subdivided accord-
ing to whether the pigment could cover large areas of the shell
or could occur only episodically as small markings. The three
episodic pigments are white, gray, and yellow. White, an
Opaque pigment distinct from the white background color
usually occurs as mottles or chevrons. Based on a
microscopic examination of the shell, this color appears

Table 1. Summary of the system used to describe and classify the
appearance of the shell.

Factor Variants

Background Color white
yellow
orange

Pattern Color white
episodic gray
yellow

slate
covering brown
chestnut

Pattern bands
episodic rays
mottle & chevron

continuous
covering ribs only

top valve only

Overall Appearance chestnut present

chestnut absent

definitely to be an applied pigment and not an absence of
pigment. The pale dove gray can merge into the white pig-
ment and can be a variant of it. This color often overlays slate
and brown. Yellow appears to be similar to background yellow,
but, unlike background, its coverage and intensity were not
uniform on a single shell. The three covering pigments, which
can occur mixed on the same shell, are: 1) slate, a dark
greeny-gray color which varied little in intensity; 2) brown, a
chocolate color ranging from pale to very dark; 3) chestnut,
an orangish to reddish brown which could vary in intensity.
Because the overall appearance of the shell was so strongly
affected by whether or not the covering pigment was chestnut,
scallops could be assigned to one of two “‘overall appearance”’
categories, ‘“‘chestnut present’ and ‘‘chestnut absent’,
regardless of other factors.

Patterns, as shown in figure 2, can be of several dif-
ferent types which are not necessarily genetically related to
one another. Like the pigments, they have been subdivided
into episodic patterns, which can occur on shells of any
background color, and overall patterns, which occur only on
white shells. Argopecten irradians irradians is usually regarded
as having a dark shell, but this is only because the com-
monest phenotype is a white shell with an overall pattern of
dark pigment.

There are three episodic patterns: ‘‘bands’’, “‘rays’’, and
“‘mottle & chevron’. “‘Bands”’ are strips of brown or chestnut
laid down parallel to the growing edge of the shell (Fig. 2a
and 2b, bottom valve; Fig. 2d, top valve). They are produced
by episodic production of pigment as the shell grows. ‘‘Rays’’
are produced by a lack of pigment along one or more of the
shell’s ribs from hinge to lip (Fig. 2e). This is a failure to pro-
duce pigments in a particular sector of the mantle edge. In
nature we have seen rays only on white shells with dark overall
patterns. However, scallops bred in a hatchery (Kraeuter

120 AMER. MALAC. BULL. 7(2) (1990)

Fig. 2. Photographs of six scallops that exemplify the various patterns described in the text. For each shell, the top (left) valve is to the left
and the bottom (right) valve, with its distinctive byssal notch, is to the right. Top and bottom valves show different patterns as follows: a) upper-
continuous with mottle, lower- continuous with bands; b) upper- continuous with chevrons, lower- continuous with mottles; c) upper- continuous,
lower- ribs only; d) upper- bands on ribs only, lower- no pigments; e) upper- continuous with rays, lower- unmarked except in juvenile region
which is continuous; f) upper- continuous, lower- no pigments (scale bars = 1 cm).

et al., 1984) have had white (presumably unpigmented) rays
on shells with orange or yellow background. ‘‘Mottle &
chevron’ are white, gray, or sometimes yellow markings that
are caused by the active, episodic secretion of a light pigment,
not by an absence of color. They occur only scattered across
one of the covering patterns. Mottles (Fig. 2a, top valve; Fig.
2b, bottom valve) are rectangular patches of pale pigment
usually limited to a section of one rib while chevrons (Fig. 2b,
top valve) are ‘‘V’-shaped and often extend across several
ribs. Whether these two have different causations is not
known, and they are grouped in this analysis as ‘‘mottle &
chevron’.

There are three covering patterns: ‘‘continuous’’, ‘‘ribs
only’, and ‘‘top valve only”’. In the ‘‘continuous’”’ pattern the
entire valve can be covered with dark pigment so that only
close examination of the auricles will reveal that the
background color is white (Fig. 2c and 2f, top valves). In ‘‘ribs
only” the continuous pigment can occur only on the tops of
the ribs and not in the furrows (Fig. 2c, bottom valve). This
gives the shell a rayed appearance that is very regular and

distinct from the episodically rayed pattern. A banded shell
can also show this pattern (Fig. 2d, top valve) because those
areas banded with dark pigment can have it on the tops of
ribs only or on both rib and furrow. In ‘‘top valve only’”’ the
continuous pigment, whether marked with an episodic pat-
tern or not, is sometimes restricted to the top valve only (Fig.
2f) or to the top valve and the juvenile portion of the bottom
valve (Fig. 2e).

FREQUENCIES OF SHELL PHENOTYPES

All the adult and juvenile scallops collected from
Lagoon and Nashaquitsa ponds were scored for appearance
of the shell using the classification system described above.
The four adult samples from each pond were first tested for
homogenity of frequencies and then pooled to estimate
phenotypic frequencies for that pond, while the juveniles from
all the spat collectors in one pond were treated as a single
sample. Three main analyses were then performed. First,
comparisons were made between the two age classes within

ELEK AND ADAMKEWICZ: SHELL COLOR IN ARGOPECTEN

Table 2. Frequencies of the three background colors on juvenile and adult shells from
Lagoon and Nashaquitsa Ponds. Using contingency chi squares, like age classes were
compared between ponds and different age classes within ponds.

Lagoon Pond Nashaquitsa Pond

121

Color Juvenile Adult Juvenile Adult
White 0.870 0.950 0.976 0.952
Yellow 0.113 0.033 0.024 0.039
Orange 0.017 0.017 0.000 0.010
N 231 302 371 310
Results of Contingency Tests (in all cases, degrees of freedom = 1):
Between ponds - Juveniles: X2 = 503 p < 0.0001
- Adults: X2 = 058p > 04 ns

Between ages - Lagoon Pond:

- Nashaquitsa Pond:

X2 = 13.15 p < 0.0003
chi X2 = 503p < 003

Table 3. Frequencies of the two types of overall appearance on juvenile and adult shells

from Lagoon and Nashaquitsa Ponds.

Lagoon Pond

Nashaquitsa Pond

Color Juvenile Adult Juvenile Adult
Chestnut Absent 0.898 0.868 0.865 0.805
Chestnut Present 0.102 0.132 0.135 0.195
N 231 302 371 310
Results of Contingency Tests (in all cases, degrees of freedom = 1):
Between ponds - Juveniles: X2 = 1.20 p > 0.25 ns
- Adults: X2 = 4.78 p < 0.03

Between ages - Lagoon Pond:

X2 = 1.00 p < 0.31 ns

- Nashaquitsa Pond: chi X? = 460p < 0.03

each pond to investigate whether frequencies changed with
age. Second, the same age classes were compared between
ponds to estimate the extent of variation between populations.
Third, the occurrences of pattern color phenotypes were
tested for positive or negative associations that might sug-
gest underlying genetic relationships.

Table 2 shows that, although yellow and orange are
definitely variants of a genetic polymorphism, their frequen-
cies were low in all groups. Frequencies of the colors in the
two adult samples were homogeneous and indistinguishable.
However, juvenile scallops differed from adults in each pond
and juveniles from Lagoon Pond had a significantly higher
proportion of yellow shells compared with juveniles from
Nashaquitsa. Results were also significant when frequencies
of the two ‘‘overall appearance”’ morphs, ‘‘chestnut present”’
and “‘chestnut absent’’ were compared (Table 3), but here the
deviant groups are adults in Nashaquitsa and juveniles in
Lagoon Pond.

Table 4 summarizes the data for four of the shell pat-
terns. Comparisons of pattern frequencies using contingen-
cy chi squares showed different results for each character.
The frequencies of shells with ‘‘ray’’ pattern were
homogeneous both between age groups and between ponds,
while the frequencies of ‘‘ribs only”’ differed significantly be-
tween age groups but not between ponds. The frequencies
of ‘‘mottle & chevron” and ‘“‘top valve only’ differed
significantly both between age groups and, for adults only,

between ponds, both being significantly more frequent among
adults in Nashaquitsa than in Lagoon Pond.

A comparison of the two ponds, by valve, for the fre-
quencies of individual pattern colors on the adult shells (Table
5) revealed more consistent differentiation between the two
ponds, with all but three of the paired comparisons (yellow
on top valves, brown and white on bottom valves) showing
significant differences between the ponds. Juveniles were not
included in this analysis because their shells might not have
had time to develop all their colors. Brown was the most com-
mon color in both ponds and chestnut the least common. Note
that all the dark covering pigments were more common on
the top than on the bottom valves, as was the episodic pig-
ment white, which was associated with the dark colors through
the pattern ‘‘mottle & chevron’. This is a quantitative expres-
sion of the qualitative observation that the top valve of adults
always appears darker than the bottom valve.

To investigate whether the occurrences of different col-
ors were associated, and, if so, whether the association was
positive or negative, the six pattern colors were tested for in-
dependence in all possible pairs using 2x2 contingency chi
squares, presence/absence of one color in the sample ver-
sus presence/absence of the other color. Only adult scallops
were included because of the possibility that juveniles had
not yet developed all of their adult coloration. For any signifi-
cant chi square, the degree of association was assessed by
the contingency coefficient, Phi = X2/N, with values ranging

122

AMER. MALAC. BULL. 7(2) (1990)

Table 4. Frequencies of the various patterns on juvenile and adult shells from Lagoon
and Nashaquitsa Ponds. For each age class in each pond, the total number of animals
counted is the same as shown in Table 3. Contingency tests were performed between
age classes in one pond and between the same ages in the two ponds with results as

discussed in the text.

Juvenile Adult
Top Bottom Top Bottom
Rays
Lagoon 0.048 -0- 0.043 0.020
Nashaquitsa 0.050 0.01 0.042 0.019
Mottle & Chevron
Lagoon 0.865 0.790 0.689 0.180
Nashaquitsa 0.897 0.850 0.858 0.300
Ribs only:
Lagoon 0.033 0.511 0.060 0.745
Nashaquitsa 0.114 0.510 0.033 0.652
Top valve only:
Lagoon — — 0.510
Nashaquitsa _— as 0.558

Table 5. Frequencies of individual pattern colors on both top and bottom valves of adult shells from Lagoon and
Nashaquitsa Ponds. Note that multiple colors can appear on one shell. In contingency comparisons of the same
valve between ponds, all comparisons were significantly different at the 0.05 level except as noted (*).

Covering Colors

Episodic Colors

Chestnut Slate Brown White Gray Yellow N
Lagoon Pond
Top 0.13 0.93 0.95 0.24 0.80 0.57* 302
Bottom 0.13 0.55 0.89* 0.05* 0.47 0.71 302
Nashaquitsa Pond
Top 0.19 0.88 0.91 0.78 0.56 0.63" 310
Bottom 0.19 0.46 0.85* 0.09" 0.35 0.80 310

from 0 (not associated) to 1 (perfectly associated). The results
(Table 6) are not shown separately for top and bottom valves
because the associations between the colors were the same
on both valves. With two exceptions (the associations between
white/brown and yellow/gray were significant in one pond but
not the other) the 15 comparisons gave the same results in
both ponds. The patterns of association, with very negative
associations of chestnut with brown, yellow, and gray and a
correspondingly strong positive association of brown with
yellow and gray, support the reality of the variants for ‘‘overall
appearance’”’ based on presence or absence of chestnut.
Strong positive associations also exist for slate with white and
slate with gray, while slate itself shows no association with
chestnut and only a modest association with brown.

Table 7 summarizes the results for shell dimensions.
Measurements of the same age classes were compared be-
tween ponds using a one way analysis of variance, and both
juveniles and adults from Lagoon Pond were significantly
larger in all linear dimensions than the same age groups from
Nashaquitsa Pond. Although relative depth was not tested for
significance, adult shells from Nashaquitsa did have more
strongly curved valves, as indicated by a greater value for
relative depth. There was no significant difference in size
associated with any pattern or color except for
presence/absence of chestnut. Shells with chestnut pigment

were on average 2 mm smaller in height and width than shells
without chestnut but were slightly deeper.

DISCUSSION

Argopecten irradians irradians exhibits a wide range
of variation in the patterns and colors of its shells. The system
proposed in this study classifies this variation into three
background colors, six pattern colors, and six patterns. These
categories have the virtue of being discrete, with only two or
three alternatives each, which leads to testable genetic
hypotheses. The significant positive and negative associations
among the six pattern colors suggest that the underlying
pigments can be a sequence of products in one, or a few,
biochemical pathways. Background color is already known
to behave as a single gene trait, and it is reasonable to believe
that the pattern colors will also be shown to be under the con-
trol of only a few genes.

The patterns themselves present a more complicated
picture, with expression influenced by both genetic and en-
vironmental factors. The genetic interaction is clearly shown
by the results of earlier breeding experiments (Adamkewicz
and Castagna, 1988) in which self-fertilized crosses of uniform-
ly orange (or yellow) scallops produced offspring of two kinds,
those with orange background (or yellow) and those with white

ELEK AND ADAMKEWICZ: SHELL COLOR IN ARGOPECTEN

background, in the standard 3:1 Mendelian ratio. Although
the parents had unpatterned valves, the patterns ‘‘mottle &
chevron’ and ‘‘top valve only”’ did appear in the offspring,
indicating recessive inheritance of these patterns. However,
the patterns occurred only on the one-quarter of the offspring
with white background color, never on the three-quarters of
the offspring with orange or yellow backgrounds. In the natural
population also, we never observed the patterns ‘‘mottle &
chevron’ or ‘‘top valve only”’ on scallops with orange or yellow
background color. Yellow and orange shells can be extensively
covered with dark pigments, but not in a ‘‘mottle & chevron’
pattern. These observations support a model in which one
or more genes determine pattern, and these gene(s) either
interact in recessive epistasis with the gene for background
color or perhaps are held in a complex supergene with the
pigment loci, as is the case in the snail Cepaea (Cain et al.,
1968). Present evidence cannot distinguish among these or
more complex models.

Another complication in the expression of pattern is
developmental and could depend on environmental in-
fluences. Regardless of background color, the top (left) valve
of a shell is always more heavily marked with pattern pigments
than is the bottom (right) valve. Furthermore, ‘‘mottle &
chevron’, the principal source of dark covering pattern, is most
common on juveniles and on the juvenile portion of adult
shells, indicating that the pattern is usually expressed on both
valves early in life but then often ceases to be produced on
the lower valve as the shell grows larger. A majority of adult
shells show the ‘‘top valve only” pattern and often, but not
always, these do have pigments in the juvenile region of the
lower valve (Fig. 2e).

Other patterns are also expressed preferentially on one
valve. The pattern ‘‘rays’’, like ‘‘mottle & chevron’, is more
common on top valves while the pattern ‘‘ribs only’ is more

123

common on bottom valves. There is very little evidence on
the genetic status of these differences in pattern. Results from
one mating suggest that the presence of the ‘‘mottle &
chevron’ pattern on both valves or on the ‘‘top valve only”’
is an allelic difference (Adamkewicz and Castagna, 1988).
However, whether one gene controls the production of
“episodic” pigments while another gene controls their
distribution in various ‘‘covering’’ patterns or whether one
complex locus with many variants controls both aspects of
patterning is completely unknown.

The possible causes of these associations between
valve, age, and pattern have never been investigated, but the
contrast between valves and the shift in the expression of
‘“‘mottle & chevron’’ do correspond to a shift in habitat that
juveniles experience. During the first months of life, juvenile
scallops cling to submerged objects, and both their valves
are exposed to light and to the view of predators. After two
or three months, the scallops drop onto the substratum and lie
on their bottom (right) valves with only the top valves exposed.
The overall lighter coloration of the lower valve may well be
a response to this change of position. The patterns cannot,
however, be entirely the product of environmental influences.
Some scallops have both valves darkly pigmented throughout
their lives, some show the ‘‘top valve only”’ pattern only on
the adult portion of the shell, and some display the ‘‘top valve
only” pattern throughout life, yet all experience the same shift
in habitat. One possible explanation is that the change of
habitat triggers a general lessening of pigmentation in the
lower valve but that the mechanism by which the lightening
is accomplished, and its extent, depends on the individual’s
particular genotype for a set of polymorphic pigment and pat-
tern loci. Such interactions of genotype, age, and environment
require that comparisons of pattern frequencies between age
groups be made with great caution.

Table 6. Associations between pairs of pattern colors on top valves of adults were investigated with
a series of 2 x 2 contingency chi squares for the presence/absence of each possible pair of colors
in each pond. The number given for each pair is the Phi value (X2/N), which measures the degree
of association. The direction of associaton is shown by use of + and — symbols while the level of
significance of the association is shown by the number + or — symbols. One, two, or three uses of
the symbol indicates significance at the 0.05, 0.01, or 0.001 level respectively while ns indicated that
the p value of the chi square was not significant. Data for Nashaquitsa Pond are in the upper right
quadrant and data for Lagoon Pond are in the lower left.

Slate Brown Chestnut White Gray Yellow
Slate ++ ns +++ ++4 ns
0.11 0.47 0.45
Brown + --- ns +++ +++
0.08 0.69 0.28 0.51
Chestnut ns --- +++ —— 2H
0.70 0.16 0.20 0.64
White +++ +44 +44 +++ ---
0.16 0.16 0.30 0.19 0.29
Gray +++ +++ --- +++ ns
0.62 0.18 0.15 0.13
Yellow ns +++ — —— ae
0.28 0.44 0.16 0.08

124 AMER. MALAC. BULL. 7(2) (1990)

Table 7. Mean dimensions in millimeters of juvenile and adult shells from Lagoon and Nashaquitsa Ponds
with standard deviations shown in parentheses. Shells from Lagoon Pond were significantly larger in all linear
dimensions than shells from Nashaquitsa, the ANOVA for each comparison having p < 0.01. Although not
tested for significance, relative depth (= depth/height) is also given.

Height Width Depth Rel. Depth N
LAGOON POND:
Juvenile mean mm 18.2 18.4 74 0.404 231
std error +034 + 0.39 +0.16
Adult mean mm 52.50 55.40 23.10 0.440 302
std error + 0.30 +033 + 0.16
NASHAQUITSA POND:
Juvenile mean mm 15.9 15.7 6.0 0.377 377
std error +013 + 0.15 + 0.06
Adult mean mm 48.20 50.70 22.10 0.456 310
std error + 0.35 + 0.37 +0.20

Regardless of how the various shell characters are pro-
duced, the significant difference in their frequencies for the
same age groups in different ponds indicates that the two
populations are relatively isolated from each other. The degree
of genetic isolation of the two pond populations was not in-
vestigated directly in this study, but observations on the move-
ment of scallop larvae out of the Niantic Estuary (Marshall,
1960; Moore and Marshall, 1967) give some indication of the
isolation to be expected. These findings, like the present ones,
suggest that there should be very little movement of larvae
between ponds on Martha's Vineyard, particularly since the
ponds open onto different bodies of water.

Differences between ponds could be a result only of
their isolation, i.e. due only to genetic drift, or a result of selec-
tion based on differences in the environments of the two
ponds. The ponds do differ in some physical parameters.
Nashaquitsa Pond is smaller in area, shallower, and takes
longer in the spring to reach the critical spawning temperature
of 20°C than does Lagoon Pond (Elek, 1985). The salinities
and substrata of the areas sampled were similar in both ponds
as were plant and algal species on the substratum, while
other aspects of the biological environment were not in-
vestigated. The earlier warming of Lagoon Pond probably ex-
plains the larger size attained by Lagoon scallops. The warmth
not only increases the growth rate but also initiates earlier
spawning, which provides a longer period for growth. As ex-
pected, if this is an important difference between the ponds,
most of the total difference in mean size between the two
populations was already achieved by the juvenile scallops.

The wide variety of colors and patterns is clearly a
polymorphism of long standing. Both Abbott (1974) and Clarke
(1965) have noted the existence of colors and patterns other
than wild-type in Argopecten. Although Clarke did not make
the distinction between background and pattern that the pre-
sent authors do, his demonstration that the frequency of white
bottom valves (the pattern ‘‘top valve only”’ of this paper)
varies geographically proves that the polymorphism for pat-
tern is widespread and of long standing. Furthermore, his
remark that only white shells occurred in frequencies high
enough to record suggests that the polymorphism is
widespread and of long standing, with yellow and orange

variants always rare. Additional evidence that the poly-
morphism for background color is neither new nor transient
comes from a private collection of shells taken from Lagoon
Pond in 1980. This sample was scored as containing 0.97
white, 0.013 yellow, and 0.017 orange shells. These counts are
very similar to those in Table 2 and suggest that the poly-
morphism for background color is a persistent genetic poly-
morphism, at least on Martha’s Vineyard. If this stability can
be confirmed, drift will be a very unlikely explanation for the
observed differences between ponds.

The highly significant difference in the frequency of
yellow between adults and juveniles of Lagoon Pond provides
some evidence that selection is acting on the polymorphism,
but its meaning is not clear. Without data from several
breeding seasons, one cannot know whether the difference
is a unique event due to chance or a regular, biologically
significant, occurrence indicating differential survival between
colors. In light of the evidence that the polymorphism is neither
new nor transient, any systematic selection against yellow at
one stage would have to be balanced by an advantage in
another area.

Those polymorphisms in marine mollusks that are, at
least in part, understood appear to be driven by response to
high temperatures. Mitton (1977) found a convincing relation-
ship between temperature and the blue/brown polymorphism
in the mussel Mytilus edulis, with the brown morph surviving
high temperatures better and being more common in the
southern part of the species range. Etter (1988) has found a
similar relationship in the snail Nucella lapillus where white
individuals survive better than brown ones in sheltered sites
where the principal stress is temperature. It seems unlikely
that temperature as a selective agent could account for dif-
ferences between these two ponds, but its role should be
carefully investigated both geographically and as one element
of possible balancing selection.

Regardless of the pond in which they occur, the fre-
quencies of the alleles for yellow and orange are low. Because
both yellow and orange alleles are dominant, their combined
frequency is estimated to be approximately 0.025 (1 - /0.95)
and, if the population is near Hardy-Weinberg equilibrium, vir-
tually all yellow and orange individuals should be heterozy-

ELEK AND ADAMKEWICZ: SHELL COLOR IN ARGOPECTEN 125

gous. The low frequencies of yellow and orange are interesting
because they are examples of a polymorphism with rare
dominants, and there has been considerable debate in the
literature over mechanisms whereby rare dominants can be
maintained in a population. Although the frequencies of yellow
and orange are low, they are still too high for a balance be-
tween recurring mutation and selection to be the likely
mechanism. Using the equation H = 2v/s to estimate selec-
tion (Falconer, 1981), selection would have to be negligible
(s<0.001) unless the mutation was very high.

Another plausible mechanism, frequency-dependent
or apostatic selection by visual predators giving an advan-
tage to any rare morph, has been proposed by Clarke (1962).
Moment (1962) has put forward a related theory of reflexive
selection, proposing that the enormous number and variety
of colors in some natural populations may be an adaptation
in itself, providing protection against predation by visually
discriminating predators such as fish and birds. Moment
(1962) cites the enormous range of variation in Donax species
as one example of reflexive selection, and Argopecten ir-
radians irradians, with its wide range of colors and patterns
may be another example of reflexive selection. Frequency
dependent selection, as described by Clarke and Moment,
could account for the persistence of rare dominants in the
stable polymorphism in the population of scallops on Martha’s
Vineyard.

The documented molluscan, echinoderm and crusta-
cean predators of scallops are unlikely to be the agents sus-
taining the polymorphism because they are primarily non-
visual predators. However, vertebrate predators such as teleost
fish and birds could provide selection pressures based on
shell color, especially in juveniles with their brighter, unfouled
colors and exposed habitat. Although there is no documen-
tation, fish are probably important predators on juvenile
scallops and could be responsible for the observed dif-
ferences between age groups and ponds. Birds have been
reported as taking quite large scallops. Gutsell (in Broom,
1976) reported that Herring Gulls, Larus argentatus Pontop-
pidan, catch many scallops at low tide, and diving ducks have
also been reported taking large numbers of shellfish. Kortright
(1967) recorded twelve species of diving ducks (sub-family:
Nyrocinae) for which mollusks comprise up to 80% of their
diet. Of these, he cited the White-Winged Scoter, Melanitta
fusca deglandi (Linnaeus), and the American Scoter, M. nigra
(Linnaeus), as major predators, with scallops forming about
half the total food of both species of scoters. Today large
numbers of diving ducks overwinter in the vicinity of Martha’s
Vineyard and are major predators of shellfish in the region.
Thus, it seems very possible that shore birds are providing
the frequency-dependent selection necessary to sustain the
polymorphism of shell color in the populations of scallops
which have been described in this paper.

Argopecten irradians has all of the elements necessary
to make it a promising subject for ecological and evolutionary
genetic studies. The species has an extensive polymorphism
with a genetic basis and with rare dominants, its populations
are known to differ in the frequencies of the morphs, and an
extensive suite of visual predators is known. In addition, the

species probably shows variation in its polymorphism on a
geographical scale. An elucidation of the forces acting on this
system would permit comparison with the polymorphisms in
other marine mollusks and an evaluation of the roles of en-
vironmental stresses such as temperature, of predation, and
of random events in maintaining natural variation.

ACKNOWLEDGMENTS

The authors are very grateful to Richard Karney for his advice
and for his assistance in making the collections. We also wish to thank
T. G. Marples and the anonymous reviewers for their comments on
this manuscript.

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126 AMER. MALAC. BULL. 7(2) (1990)

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Date of manuscript acceptance: 27 November 1989.

PREHISTORIC FRESHWATER MUSSEL (NAIAD) ASSEMBLAGES
FROM SOUTHWESTERN IOWA

JAMES L. THELER
DEPARTMENT OF SOCIOLOGY AND ANTHROPOLOGY
UNIVERSITY OF WISCONSIN, LA CROSSE
LA CROSSE, WISCONSIN 54601, U.S.A.

ABSTRACT

Archaeological excavations at nine prehistoric Indian sites along the Missouri River drainage
of southwestern lowa produced 275 freshwater mussel (naiad) valves representing at least 13 species.
These subfossils are the remains of mollusks collected as a food resource and to obtain shells for
use as tools. While little historic data are available for this region, distribution records show that all
13 mussel taxa were widespread historically in the south central Missouri River drainage. The arch-
aeological data indicate that molluscan communities having a similar species composition to those
in the region today have been present for a millennium or more.

There is no published information on the distribution
of freshwater mussel (Mollusca: Bivalvia: Unionidae) species
in the streams and rivers of southwestern lowa. It can be
assumed from historic distributional data that the streams of
this area once supported a mussel fauna similar to that of
adjacent regions. The freshwater mussel valves recovered
from prehistoric Indian sites in southwestern lowa provide an
opportunity to evaluate the distribution of naiad mollusks of
this region prior to EuroAmerican settlement.

Mussel valves from nine archaeological sites in
southwestern lowa are considered in this report. The oldest
mussel assemblage is from the Hanging Valley Site (13HR28),
which overlooks the Little Sioux River near its confluence with
the Missouri River in Harrison County, lowa (Fig. 1). Hang-
ing Valley is a Woodland Tradition cemetery and habitation
site used at about A.D. 400 (Tiffany et a/., 1988). The remain-
ing eight archaeological sites are Glenwood Culture earth-
lodges, occupied between approximately A.D. 1000 and A.D.
1300 (Hotopp, 1978). The Glenwood earthlodges are located
in Mills County, lowa; seven of these were situated along the
Pony and Keg creeks which form a small tributary to the
Missouri River. The other earthlodge (13ML176) was located
immediately adjacent to the Missouri River valley a few miles
north of the Pony and Keg creeks cluster (Fig. 1).

METHODS AND MATERIALS

The southwestern lowa subfossil mussel valves were
identified by the author at the University of Wisconsin, La
Crosse, and now are housed permanently in the Archaeolog-

Little
Sioux
River

Tarkio River

ML176 wep (2 branches)

Keg& Pony
Creek Sites

Platte
River

Fig. 1. Location of archaeological sites in southwestern lowa from
which freshwater mussel valves were obtained.

ical Repository at the Office of the State Archaeologist, Univer-
sity of lowa, lowa City, lowa. Naiad taxonomy used in this
report follows the nomenclature presented by Turgeon et al.
(1988). Table 1 provides a listing of the number of valves of
each taxon for each archaeological site. A comparison of the
subfossil naiades with historic naiad faunas recorded from
the Missouri River and its tributary streams in northwestern
and west central lowa, northwest Missouri and portions of
North and South Dakota are presented in Table 2.

RESULTS

In all, 275 valves of 13 mussel species are represented

American Malacological Bulletin, Vol. 7(2) (1990):127-130

127

128 AMER. MALAC. BULL. 7(2) (1990)

Table 1. Distribution and number of freshwater mussel valves recovered from prehistoric archaeological sites in southwestern lowa.

Little Missouri
Sioux R. River

Drainage:

Site Number: 13HR28 ML176

Keg Creek

ML128 ML130 ML131

Pony Creek

Total %

ML132 ML135 ML126 ML136 Valves of Total

Family Unionidae
Subfamily Anodontinae

Anodonta grandis _ 1 = mae
Lasmigona complanata
complanata — 6 = =

Subfamily Ambleminae
Tritogonia verrucosa 2
Quadrula quadrula —
Q. pustulosa pustulosa — 1
Amblema plicata plicata — 45
Fusconaia flava — 6

hm Pw

| w |
= |

Subfamily Lampsilinae
Leptodea fragilis 2 —
Potamilus alatus 18 1
Ligumia recta 4
Lampsilis teres 1 3
L. siliquoidea — 4 — —
L. cardium 1 22
L. sp. _ _

Totals 24 99 3 ih

— — 1 — — 2 0.7
1 2 1 — 3 13 4.7
_ — 1 1 3 11 40
— — 1 1 — 8 29
—_ _ _— = = 1 0.4
4 8 8 7 10 86 31.3
_ 3 1 3 1 14 54
1 — — = - 3 A
3 1 1 1 5 30 10.9
1 2 3 — 4 14 51
_ ~— — = — 4 15
—_ — 3 — — rf 25
12 14 21 6 79 28.7
— — — 1 2 3 11
22 30 4 15 34 275 100.0

at the nine southwestern lowa archaeological sites (Table 1).
The two most abundant species were Amblema plicata plicata
(Say, 1817), with 86 valves equalling 31.3% of the combined
site specimens, and Lampsilis cardium (Rafinesque, 1820),
with 79 valves (28.7%); shells of both taxa occurred at eight
of the nine states. Potamilus alatus (Say, 1817) recovered at
seven sites was next in abundance with 30 valves represent-
ing 10.9% of all naiad valves. Fusconaia flava (Rafinesque,
1820) and Ligumia recta (Lamarck, 1819), both having a total
of 14 valves each (5.1%), were recovered at five of the nine
sites and rank fourth in abundance. The remaining eight taxa,
in decreasing frequency of occurrence, are given in Table 1.

DISCUSSION

Utterback (1915) presented the distribution of mussel
taxa found in three small northwest Missouri rivers that flow
from southwestern lowa. These include, from east to west,
the Platte, Nodoway and Tarkio rivers, but unfortunately, the
Nishnabotna River, flowing almost entirely through south-
western lowa, was not considered. All of the above streams
drain into the Missouri River. The distribution of freshwater
mussels in the Missouri River along the Nebraska border with
lowa was presented by Hoke (1983). In west central lowa, data
on the modern distribution of mussels in the middle and up-
per portions of the Little Sioux River were presented by
Rausch and Bovbjerg (1973). The naiad fauna of other
Missouri tributaries including the Big Sioux, James and Ver-
million rivers in northwest lowa, and the Dakotas was reported

by Coker and Southall (1914) (Figs. 1, 2).

A comparison of the southwestern lowa subfossil
mussel assemblage composition with survey data for streams
in the adjacent parts of the Missouri River basin shows that
all 13 subfossil taxa were recorded historically in the region.
Eleven species, Anodonta grandis, Lasmigona complanata
complanata, Tritogonia verrucosa, Quadrula quadrula, Q.
pustulosa pustulosa, Amblema plicata plicata, Fusconaia flava,
Leptodea fragilis, Potamilus alatus, Ligumia recta and Lamp-
silis teres are recorded for northwestern Missouri streams
draining southwestern lowa, while two species represented
in the archaeological assemblage, Lampsilis siliquoidea and
L. cardium, are not recorded historically from the Missouri
River adjacent to lowa (Hoke, 1983) or from northwestern
Missouri streams (Utterback, 1915-16; Oesch, 1984). However,
both L. cardium and L. siliquoidea have been recorded in
historic times in the Missouri River drainage north of south-
western lowa (Table 2).

Although valves representing many mussel taxa
recovered at the southwestern lowa archaeological sites ex-
hibited signs of human modification by grinding and/or flak-
ing to produce tools, the large cup-like valves of Lampsilis
cardium seem to have been especially favored by prehistoric
peoples who easily converted these shells into a variety of
spoons and cups. It is probable that L. cardium valves were
intentionally sought by aboriginal peoples for utensil produc-
tion. This taxon’s widespread occurrence and the presence
of numbers of unmodified valves seems to argue for local
populations in suitable southwestern lowa habitat prior to

THELER: PREHISTORIC NAIAD FAUNA 129

EuroAmerican settlement.

Six naiad taxa having a limited historic distribution in
the Missouri River drainage are absent from the southwestern
lowa archaeological record; these include Anodonta sub-
orbiculata Say, 1831, Strophitus undulatus (Say, 1817),
Alasmidonta marginata Say, 1818, Arcidens confragosus (Say,
1829), Obliquaria reflexa Rafinesque, 1820 and Truncilla
donaciformis (Lea, 1828). Potamilus ohiensis (Rafinesque,
1820), while widespread historically in this portion of the
Missouri River basin (Table 2), also is not present in the
southwest lowa subfossil assemblage. Potamilus ohiensis is
at the northwestern margin of its range in the Missouri River
drainage and has perhaps occupied (or at least become
widespread in) this region only in the historic period.
Pleurobema coccineum (Conrad, 1834) is recorded during
historic times in at least three streams feeding the Missouri
River, but has not been recorded as a southwestern lowa
subfossil.

Freshwater mussel valves also have been reported in
prehistoric context at the Woodland Tradition Rainbow site
(18PM91) in Plymouth County, lowa (Riggle and Freitag, 1981).
This northwestern lowa site is located adjacent to the Floyd
River, a tributary to the Missouri River. The Rainbow site naiad

Fig. 2. Location of selected streams with documented freshwater
mussel populations in the south central Missouri River drainage.

Table 2. A comparison of naiad shells from archaeological sites in southwestern lowa with historic naiad distribution in portions of the Missouri
River drainage (P, taxon present; —, taxon absent) (1, Hoke, 1983; 2, Utterback, 1915-16; 3, Rausch and Bovbjerg, 1973; 4, Coker and Southall, 1914).

Data Source: This Report

Subfossils Missouri R.1 NW
(n=275 Adjacent to Missouri? Little Big Ver-
valves) lowa Tarkio R. Nodaway R. Platte R. Sioux R.2 Sioux R.4 James R.4 million R.4
Family Unionidae
Subfamily Anodontinae
Anodonta suborbiculata _— P — — — = = aes =
A. grandis 0.7 P P = P P _ P P
Strophitus undulatus — — — = P = — <= =
Alasmidonta marginata = _— — — — = P = =
Arcidens confragosus —_— — — — = _ — P —_
Lasmigona complanata
complanata 4.7 P P P P P P P P
Subfamily Ambleminae
Tritogonia verrucosa 4.0 —_ P P P — — P =
Quadrula quadrula 29 — P P P P — P P
Q. pustulosa pustulosa 0.4 — P P P = P Pp P
Amblema plicata plicata 31.3 _— P P P P P P P
Fusconaia flava 51 = = — >) = = es _
Pleurobema coccineum — — — = = — P P
Subfamily Lampsilinae
Obliquaria reflexa _ — = = P = = — as
Truncilla donaciformis — — = = Pp — == _ =
Leptodea fragilis 11 P P P P P >) Pp )
Potamilus alatus 10.9 — P P P — P P Pp
P. ohiensis — P P P P P = = za
Ligumia recta 5.1 — _— = P P P P )
Lampsilis teres 15 — P — P = = P =
L. siliquoidea 25 = — — — _— P P P
L. cardium 28.7 _ — — — P P P

L. sp. 1.1 = ee

|
|
|
|
|
|

130 AMER. MALAC. BULL. 7(2) (1990)

assemblage contains 11 of the 13 taxa recorded as subfossils
in southwestern lowa, lacking only Tritogonia verrucosa and
Quaarula quadrula. Three species, Lasmigona costata
(Rafinesque, 1820), L. compressa (Lea, 1829) and Actinonaias
ligamentina (Lamarck, 1819) not recovered in the southwestern
lowa subfossil assemblages or recorded in this region dur-
ing the historic period, are reported as members of the Rain-
bow site subfossil assemblage. All three of the above taxa
are on their range margin in the Missouri River drainage of
northwestern lowa, judging from historic distributions (Clarke,
1985; La Rocque, 1967). Like the southwestern lowa arch-
aeological assemblages, the Rainbow subfossil material did
not contain Anodonta suborbiculata or Potamilus ohiensis.

The distribution of common and rare mussel taxa found
in southwestern lowa is paralleled in other aboriginal
assemblages when compared to adjacent modern stream
faunas. Klippel et a/. (1978) reported 25 species of freshwater
mussel from the modern Pomme de Terre River of western
Missouri, while archaeological deposits at nearby Rodgers
Shelter contained only 16 species. The authors indicate that
10 of the 11 species absent from Rodgers Shelter are small
size or low frequency taxa that could have failed to have been
collected by aboriginal mussel harvestors. Thin shelled taxa,
missing from the assemblage, may not have been preserved
Klippel et a/. (1978).

The Mississippi River in the vicinity of Prairie du Chien,
Wisconsin, once contained approximately 44 mussel species
(Havlik and Stansbery, 1978). Extensive aboriginal shell
deposits at Prairie du Chien were found to contain 28 mussel
species (Theler, 1987). As is the case in southwestern lowa,
reported here, and western Missouri (Klippel et a/., 1978),
those species absent from archaeological deposits are, for
the most part, taxa that are of small size or are those found
to be rare in historic times (Theler, 1987). As with all archaeo-
logical assemblages mussels, those of southwestern lowa
have passed through a ‘‘filter’’ of human cultural behavior
and must be evaluated in that light.

CONCLUSIONS

The assemblages of subfossil mussel valves recovered
from nine archaeological sites in southwestern lowa contained
13 of 21 taxa recorded during the historic period for the south
central portion of the Missouri River and its tributaries con-
sidered in this report. The 13 species represented as
archaeological subfossils were found to be widespread in the
region historically. Generally, those naiad taxa that were rare
in historic surveys were species not represented among the
southwest lowa archaeological assemblages. One exception
is Potamilus ohiensis, found to be widespread in the region
during the historic period, but absent from the subfossil
assemblages. It is possible that P ohiensis could have extend-
ed its range or increased in abundance in the Missouri River
basin since EuroAmerican settlement of the region.

ACKNOWLEDGMENTS

| would like to thank the former State Archaeologist of lowa,
Duane Anderson, and his associates at the University of lowa for their

assistance during this study. | am particularly grateful to William
Billeck, LuAnn Hudson, Kris Hirst, Joseph Tiffany, and William Green
for their assistance. Susann Theler and Teri Stueck are acknowledged
for typing drafts of this paper. | would like to thank the three
anonymous reviewers for their insightful comments on earlier drafts
of this report.

LITERATURE CITED

Clarke, Arthur H. 1985. The Tribe Alasmidontini (Unionidae:
Anodontinae), Part Il: Lasmigona and Simpsonaias. Smith-
sonian Contributions to Zoology, No. 399, 75 pp.

Coker, Robert E. and John B. Southall. 1914. Mussel Resources in
Tributaries of the Upper Missouri River. Report of the United
States Commissioner of Fisheries for 1914, Appendix 4:1-17
(issued separately as Bureau of Fisheries Document No. 812).

Havlik, M. E. and D. H. Stansbery. 1978. The Naiad Mollusks of the
Mississippi River in the Vicinity of Prairie du Chien, Wiscon-
sin. Bulletin of the American Malacological Union for 1977:9-12.

Hoke, Ellet. 1983. Unionid Mollusks of the Missouri River on the
Nebraska Border. American Malacological Bulletin 1:71-74.

Hotopp, John. 1978. Glenwood: A Contemporary View. In: The Cen-
tral Plains Tradition: Internal Development and External Rela-
tionships, Donald J. Blakeslee, ed. pp. 109-133. Report No. 11,
Office of the State Archaeologist, The University of lowa (lowa
City).

Klippel, W. E., G. Celmer, and J. R. Purdue. 1978. The Holocene Naiad
Record at Rodgers Shelter in the Western Ozark Highland of
Missouri. Plains Anthropologist 23(82):257-271.

La Rocque, Aurele. 1967. Pleistocene Mollusca of Ohio, Part 2. Ohio
Department of Natural Resources, Division of Geological
Survey, Bulletin No. 62.

Oesch, Ronald D. 1984. Missouri Naiades: A Guide to the Mussels
of Missouri. Missouri Department of Conservation, Jefferson
City, Missouri. 270 pp.

Rausch, Clair G. and Richard V. Bovbjerg. 1973. Fauna of the Mid-
dle Little Sioux River and Comparison with Upper and Lower
Regions. Proceedings of the lowa Academy of Science
80:111-116.

Riggle, R. Stanley and Thomas M. Freitag. 1981. The Fresh-water
Mussels (Mollusca: Bivalvia: Unionidae) of the Rainbow Site:
In: Archaeological Investigations at the Rainbow Site, Plymouth
County, lowa, David W. Benn, ed. pp. 282-302. Contract No.
C 3571 (78). Interagency Archaeological Services, Denver.

Theler, James L. 1987. Prehistoric Freshwater Mussel Assemblages
of the Mississippi River in Southwestern Wisconsin. Nautilus
101(3):143-150.

Tiffany, Joseph A., S. J. Schermer, J. L. Theler, D. W. Owsley, D. C.
Anderson, E. A. Bettis and D. M. Thompson. 1988. The Hang-
ing Valley Site (13HR28): A Stratified Woodland Burial Locale
in Western lowa. Plains Anthropologist 33:219-257.

Turgeon, D. D., A. E. Bogan, E. V. Coan, W. K. Emerson, W. G. Lyons,
W. L. Pratt, C. F. E. Roper, A. Scheltema, F. G. Thompson,
and J. D. Williams. 1988. Common and scientific names of
aquatic invertebrates from the United States and Canada:
mollusks. American Fisheries Society Special Publication No.
16, 277 pp.

Utterback, William |. 1915. Naiadgeography of Missouri. American
Midland Naturalist 4:26-30.

Utterback, William |. 1915-1916. The Naiades of Missouri. American
Midland Naturalist 4:41-53, 97-152, 181-204, 244-273, 311-327,
339-354, 387-400, 432-464.

Date of manuscript acceptance: 9 August 1989.

RESEARCH NOTE

RECTIFICATION OF THE NOMENCLATURE
OF CERTAIN SPECIES OF TRICULINE SNAILS TRANSMITTING
PARAGONIMUS AND SCHISTOSOMA IN CHINA

LIU YUE YING
INSTITUTE OF ZOOLOGY
ACADEMIA SINICA
BEIJING, CHINA

GEORGE M. DAVIS
ACADEMY OF NATURAL SCIENCES
OF PHILADELPHIA
PHILADELPHIA, PENNSYLVANIA 19103, U.S.A.

To date, 12 species that transmit Paragonimus or
Schistosoma in China have been classified as Tricula Ben-
son, 1843 (Liu, ef a/., 1980, 1983a, b, 1984; Liu, 1983 and un-
pub. data). Tricula has been considered by Chinese
workers to belong to the family Hydrobiidae (Liu et a/. 1980,
1983a, b; Kang, 1983, 1984; Liu, 1983). However, intensive
studies of Tricula and allied genera have shown that the family
Hydrobiidae does not occur in China or Southeast Asia; Tricula
belongs to the Pomatiopsidae; Triculinae: Triculini (Davis, 1979,
1980; Davis et a/. 1983, 1984, 1986a, b).

Recent collaboration between the Institute of Zoology
and the Academy of Natural Sciences has enabled us to re-
examine all Triculinae species names associated with the
transmission of human disease. This was necessary because
early identifications were based on the original descriptions
of Gredler (1885-1892), Heude (1890) and Annandale (1924a,
b) and the photographs of Gredler’s types by Yen (1939). All
of these sources were truly inadequate in detail of descrip-
tion, size of printed photographs, and illustrations showing
intrapopulation variation and accordingly misidentifications
were inevitable. One of us (Davis) has recently studied and
photographed the type specimens and it is now possible to
reevaluate the identifications of specimens in China
associated with disease transmission. Annandale’s types are
in the British Museum; Gredler’s paralectotypes are in the
Senckenberg Museum, Frankfurt am Main, Germany;
Heude’s Tricula types are in the Museum of Comparative
Zoology at Harvard University and the Academy of Natural
Sciences of Philadelphia.

A complicating factor is that species currently assigned
to Tricula in China actually belong to at least two genera:

Tricula Benson, 1843 and Neotricula Davis 1986. Generic
distinction is based on detailed comparative anatomy; one
cannot separate the genera on shell or radular characters.
The generic distinctions are illustrated in figure 1. The descrip-
tion of Tricula is based on the comparative anatomy of north-
ern India 7. montana Benson, 1843; the type species is from
northern India (Davis et a/., 1986b). The description of
Neotricula is based on Lithoglyphopsis aperta Temcharoen
1971 [a species later relegated to Tricula (Davis, 1979) from
the Mekong River of Thailand and Laos].

Neotricula: 1) The oviduct runs from gonad to the pallial
oviduct without making a twist or coil; 2) the duct of the
seminal receptacle arises from the duct of the bursa (or
spermathecal duct); 3) the spermathecal duct runs to the end
of the mantle cavity beside the pericardium; it does not enter
the pericardium; 4) a slender sperm duct connects the duct
of the bursa to the oviduct; 5) the wall of the pericardium is
a thin unspecialized membrane; 6) The pericardium does not
bulge out into the mantle cavity.

Tricula: 1) The oviduct makes a tight coil or twist dor-
sal to the bursa copulatrix; 2) the duct of the seminal recep-
tacle arises from the oviduct; 3) the spermathecal duct runs
to, and enters the pericardium; 4) there is no sperm duct; the
duct of the bursa joins the oviduct; 5) the wall of the pericar-
dium is considerably thickened and specialized to accom-
modate sperm; 6) the pericardium bulges out into the man-
tle cavity.

A second complicating factor is that there are
numerous species of Tricula sensu lato (Tricula as understood
prior to Davis et al., 1986b) spread throughout southern China.
There are at least 20 valid species known today. There are

American Malacological Bulletin, Vol. 7(2) (1990):131-133

131

132 AMER. MALAC. BULL. 7(2) (1990)

A Bu
Dbu

Opo
Dsr

Sd
Sr—___ u

>
0 eee y

Emc
oY Pe
Bu
B Dbu
Opo
Sr
Ocoi whe Dsr
Emc
Ov
Pe

Sd

Fig. 1. Bursa copulatrix complex of organs showing differences
between Neotricula (A) and Tricu/a (B) [Bu, bursa copulatrix; Dbu,
duct of the bursa; Dsr, duct of the seminal receptacle; Emc, posterior
end of the mantle cavity; Ocoi, oviduct coil; Opo, opening of pallial
oviduct into the albumen gland (posterior pallial oviduct); Ov, oviduct;
Pe, pericardium; Sd, spermathecal duct; Sdu, sperm duct; Sr, seminal
receptacle).

many undescribed species; the number of newly described
species increases each year and will continue to do so for
some time. We estimate that there are more than 60 such
species throughout China (see Kang, 1983, 1984a, b, 1986;
Liu et a/., 1983a, b, 1987; Guo and Gu, 1985; Davis, et a/.,
1986a).

A third complicating factor is that confirmation that a
species transmits a parasite is dependent on voucher
specimens cataloged into museum or institutional collections.
The voucher specimen system has not been used in China
except for type specimens; we encourage its use. The number
assigned to the specimens should be referenced in the
publication linking a parasite to the snail population in ques-
tion. This assures future investigators that the specimens seen
in acollection are the ones specified in the publication. A poor
illustration of a single specimen in a publication does not serve
to identify the species.

Given the above difficulties, we comment on eight taxa
currently listed in the literature as transmitting parasites for
which nomenclatural changes are necessary or where there

is substantial confusion. We have anatomical data for only
three of these, i.e. Neotricula jinhongensis, Tricula gregoriana,
and T. montana. All others must be retained in T sensu lato
until anatomical data are available.

1. Tricula guangxiensis Liu et al., 1983a. This is a
synonym of 7 fuchsi (Gredler 1887). 7. fuchsi transmits
Paragonimus skrjabini.

2. Tricula minutoides (Gredler, 1885): Specimens from
Hunan [Institute of Zoology, Academia Sinica, (IZAS) catalog
number 00643] are actually 7. cristella Gredler 1887. T. cristella
transmits Paragonimus skrjabini.

3. Tricula cristella (Gredler, 1887): (IZAS No. 00615):
These specimens are not that species; they belong to an ap-
parently undescribed species. Genuine T. cristella does not
transmit Paragonimus hueitungensis.

4. Tricula gregoriana Annandale, 1924a: There is much
confusion in the literature concerning this species. Specimens
figured by Liu et al. (1984) (IZAS No. 00642) are not that
species but Delavaya dianchiensis Davis and Guo, 1986 (in
Davis et al/., 1986a). Illustrations published by Sun (1959) as
T. gregoriana are also not that species. Davis et a/. (1986a)
published a description of the anatomy of snails from Yun-
nan referable to 7. gregoriana by comparison with the types.
There specimens are deposited in the collections of the In-
stitute of Zoology, Beijing with IZAS No. 08701 - (F).

5. Tricula humida Heude, 1890: There is much confu-
sion concerning the identity of this species. It is possible that
specimens illustrated by Sun (1959) are this species, but it
is not possible to confirm the identification from the reduced
photographs. We have not seen any specimens in various col-
lections in China that compare favorably with the type series.
Accordingly, we cannot confirm that 7) humida transmits
Paragonimus.

6. Tricula jinhongensis Guo and Gu, 1985: This is a
species of Neotricula; it transmits Paragonimus skrjabini and
Schistosoma sp. The parasite has not been identified to
species.

7. Tricula montana Benson, 1843. Contrary to Liu et al.,
1983a, this species does not occur in China. It is restricted
to the lesser Himalayan mountains of northern India west of
Nepal. On the basis of comparative anatomy the closest
association with Tricula in China is with 7. gregoriana.

8. Halewisia sinica Liu et al., 1983b: The genus
Halewisia is confined to the lower Mekong River in Thailand,
Laos and Cambodia. H. sinica from China is possibly Tricula
sensu lato but generic confirmation will depend on anatomical
data. T. sinica transmits Paragonimus skrjabini.

In conclusion, it is clear that considerable confusion
would be avoided if specimens found to transmit parasites
were formally cataloged into a permanent institutional collec-
tion with samples also deposited in various national centers.
The catalog numbers should be published with the data.
Descriptions of species and assignment to genera demands
detailed comparative anatomical data; it is no longer accept-
able to describe species of Triculinae on the standards of
Stimpson (1865): shell, radula, operculum, penis. Characters
of these structures are too convergent to use for these
purposes.

LIU AND DAVIS: TRICULINE SNAILS 133

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Davis, G. M., Y. H. Kuo, K. E. Hoagland, P. L. Chen, H. M. Yang,
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Date of manuscript acceptance: 2 March 1989.

SYMPOSIUM ON THE BIOLOGY
OF THE SCAPHOPODA

ORGANIZED BY
RONALD L. SHIMEK
PARAMETRIX, INC.

AMERICAN MALACOLOGICAL UNION

LOS ANGELES, CALIFORNIA
27 JUNE 1989

135

FUNCTIONAL MORPHOLOGY OF THE PERIANAL SINUS AND
PERICARDIUM OF DENTALIUM RECTIUS (MOLLUSCA: SCAPHOPODA)
WITH A REINTERPRETATION OF THE SCAPHOPOD HEART

PATRICK D. REYNOLDS
BIOLOGY DEPARTMENT, UNIVERSITY OF VICTORIA,
VICTORIA, BRITISH COLUMBIA, V8W 2Y2, CANADA

ABSTRACT

The anatomy and ultrastructure of the perianal blood sinus and pericardium in the scaphopod
Dentalium rectius Carpenter were investigated. The perianal blood sinus surrounds the rectum and
lies adjacent to the anterior wall of the pericardial coelom; the sinus is enclosed by smooth musculature
with additional muscular trabeculae traversing the sinus. The pericardium is contractile and consists
of a simple, flat epithelium with interspersed muscle fibres; both are separated from the haemocoel
by a basal lamina. The pericardial musculature is arranged as laterally oriented trabeculae which pro-
duce localized transverse constrictions of the dorsal pericardial wall. There is no evidence for a heart
enclosed by the dorsal wall of the pericardial coelom in a position ventral to the stomach as inter-
preted by earlier workers, as both a myocardium and distinct epicardium are absent. A portion of the
pericardial epithelium apposing the perianal sinus musculature is developed into podocytes and could
be the site of primary urine production. Although organogenetic information on scaphopod coelom
formation is lacking, structural similarities of the perianal sinus and pericardium in D. rectius to the

heart and pericardium in other molluscan classes support the homology of these organs.

The morphology of scaphopod circulatory structures
received a great deal of attention up to the early part of this
century, producing several conflicting interpretations of struc-
ture and function. Deshayes’ (1825) and Clark’s (1849) descrip-
tions of a heart in Dentalium spp. appear to be mistaken iden-
tifications of the esophagus and stomach, respectively.
Lacaze-Duthiers (1857) found no structure analogous to a
molluscan heart in Dentalium sp., i.e. a pulsatile vessel within
a pericardium responsible for the movement of blood, and
he considered the contractions of the pedal, perianal and ab-
dominal blood sinuses to be sufficient for circulation. A small
serous Sac within the anterior abdominal sinus, and lying be-
tween the stomach and ventral body wall, was suggested by
Lacaze-Duthiers (1857) as the pericardial rudiment; he also
noted the structural similarities of the perianal sinus to the
bivalve ventricle. Fol (1889), studying D. entalis (Deshayes),
concluded that the perianal sinus is homologous with the
heart of other molluscs.

Plate (1891, 1892) described a completely different
structure as representing the scaphopod heart; an invagina-
tion of the dorsal pericardial wall, lying ventral to the stomach
and within the pericardial coelom. Boissevain (1904) and
Distaso (1905) confirmed these results and agreed with this
interpretation. While Potts (1967) states that the pericardium
is absent in scaphopods and the heart is represented by a

contractile vessel, most recent reviews accept Plate’s findings,
at least tentatively (Fischer-Piette and Franc, 1968; Martin,
1983; Andrews, 1988).

Defining the structure of the scaphopod heart and
pericardium accurately and conclusively is of importance not
only in ascertaining the level of organization of the circulatory
system, but also in determining the role of the heart in excre-
tion or, alternatively, the structural modification of the excretory
system in the absence of a functional heart in this molluscan
class. The general organization of the excretory system in the
Mollusca is based on a haemocoel-pericardium-kidney com-
plex. Physiological work on prosobranchs (Harrison, 1962;
Little, 1965), coleoid cephalopods (Harrison and Martin, 1965;
Martin and Aldrich, 1970) and bivalves (Jones and Peggs,
1983; Hevert, 1984) has established that primary urine is pro-
duced by ultrafiltration into the pericardial coelom. The site
of ultrafiltration, as characterized ultrastructurally by the
presence of podocytes, varies from the auricular or ventricular
wall in prosobranch gastropods (Andrews, 1981),
polyplacophorans (@kland, 1980), protobranch and
pteriomorph bivalves (Pirie and George, 1979; Meyhofer et
al., 1985), to the branchial heart wall in cephalopods (Witmer
and Martin, 1973; Schipp and Hevert, 1981) and the antero-
dorsal wall of the pericardium in heterodont bivalves (Meyhofer
et al., 1985). In all cases, the ultrafiltrate enters the pericardial

American Malacological Bulletin, Vol. 7(2) (1990):137-146

137

138 AMER. MALAC. BULL. 7(2) (1990)

cavity from the haemocoel and passes through a renoperi-
cardial connection to the kidney lumen. Further modification
of the primary urine by reabsorption and secretion takes place
before excretion to the external environment via the mantle
Cavity.

Localization of an ultrafiltration barrier by ammoniacal
carmine injection, a technique used extensively in the study
of circulation and excretion up to the early 1900’s, has been
attempted in representatives of most molluscan classes and
has served as a useful basis for subsequent morphological
and quantitative physiological investigation in many
representative species (for a review, see Martin, 1983). In
scaphopods, however, it is the only physiological method ap-
plied to date, and possible sites of ultrafiltration have not been
clearly indicated. Working with Dentalium sp., Kowalevsky
(1889) noted the accumulation of ammoniacal carmine in
unspecified blood spaces and connective tissue cells. Using
the same method with D. vulgare Da Costa, Cuénot (1899)
found that these cells contain yellowish, oily granules and
broadly described their distribution as similar to that in
amphineurans and gastropods, being found under the
epithelium, between the viscera and within the interstices of
muscle fibres. On the basis of an uncertain, but presumed
common excretory function, Cuénot (1899) aligned these am-
moniacal carmine accumulating cells and those of amphi-
neurans and gastropods with the pericardial glands of bivalves
and branchial hearts of cephalopods. Strohl (1924) agreed,
labelling the cells collectively as carmine athrocytes.

An internal opening between the paired kidneys and
another coelomic (pericardial) space is absent in Dentalium
sp. according to Lacaze-Duthiers (1857), Fol (1885, 1889) and
Plate (1888, 1892). Of those investigations which describe a
pericardium, only Distaso (1905) noted a morphological con-
nection represented by a pore leading to the left kidney.

The present study of Dentalium rectius Carpenter
(Order Dentaliida) aims to clarify the morphological relation-
ship between the perianal and abdominal haemal sinuses,
the pericardial coelom and the kidney using light and
electron microscopy. The tissues of the pericardium and
associated blood sinuses are described ultrastructurally with
particular reference to contractile elements, and with a view
to identifying possible sites for ultrafiltration of blood. The in-
formation contributes toward a better understanding of cir-
culation and excretion in the Scaphopoda, and relationships
of the class within the Mollusca.

MATERIALS AND METHODS

Specimens of Dentalium rectius were dredged from ap-
proximately 60 m at Satellite Channel, close to Victoria, British
Columbia, Canada. For anatomical examination at the
light microscope level, tissues were fixed in 10% seawater--
buffered formalin, dehydrated in a graded ethanol series and
embedded in paraffin. Serial sections of 6-8 um thickness
were stained with eriochrome cyanin (Chapman, 1977).
Additionally, serial 1 wm sections of resin embedded material,
prepared as described below for transmission electron

microscopy (TEM), were stained with methylene blue-
azure Il.

Tissues for electron microscopical examination were
dissected from specimens and fixed in 2.5% glutaraldehyde
in 0.2M phosphate buffer (pH 7.4) and 0.14M NaCl for 2 hours
at room temperature. After rinsing in 0.2M phosphate buffer
and 0.3M NaCl, they were post-fixed using 1% osmium tetrox-
ide in 0.1M phosphate buffer and 0.375M NaCl for one hour
at 4°C. Tissues were rinsed in distilled water and dehydrated
in a graded series of ethanol. Specimens for scanning elec-
tron microscopy (SEM) were critical point dried from COs,
sputter coated with gold and examined in a JEOL JSM-35.
Specimens for TEM were transferred to propylene oxide
before embedding in epon resin. Ultrathin sections (grey-
silver-pale gold interference colour) were obtained on a
Reichert ultramicrotome and stained with uranyl acetate and
lead citrate (Reynolds, 1963) prior to viewing in a Philips
EM-300 transmission electron microscope.

Observations of live animals were made using a Wild
M5A dissecting microscope. Removal of the shell and a ven-
tral dissection of the mantle wall revealed the anus and
transparent ventral body wall through which the pericardium
could be seen easily.

RESULTS

PERIANAL SINUS

The perianal sinus surrounds the rectum as it passes
between the kidneys to the mantle cavity. The sinus is posi-
tioned antero-ventrally to the kidneys and the pericardium,
and no other coelomic space surrounds or apposes it (Fig.
1). The sinus is surrounded by circular and longitudinal
musculature, is traversed by an array of muscle fibres or
trabeculae, and has additional longitudinal and circular fibres
along the inner wall of the sinus enveloping the rectum (Fig. 2).

The musculature of the perianal sinus is smooth.
Neural processes occur adjacent to muscle cells, although
no synapses have been observed (Fig. 3). The muscle cells
contain thick (29-58 nm diameter) and thin (5.8 nm diameter)
myofilaments which are interspersed with a-glycogen gran-
ules (17.4 nm). Similar granules are also found concen-
trated at the periphery of the cell adjacent to the 6-11 nm wide
sarcolemma (Fig. 4). Thick myofilaments have an axial
periodicity of 9-15 nm (Fig. 5). Mitochondria are located in
clusters within sarcoplasmic bulges adjacent to contractile
elements (Fig. 4).

The muscular walls of the perianal sinus repeatedly
contract, causing an extension of the rectum and closing of
the anus, followed by relaxation of the rectum and dilation
of the anus, occurring at a rate of approximately 40-60 con-
tractions per minute. Occasional periods of relaxed dilation
last from 10 to 30 seconds. These contractions, in addition
to propelling blood through the sinus, appear to facilitate the
voiding of strings of faecal material from the rectum.

PERICARDIUM AND DORSAL
PERICARDIAL FOLDS

The pericardial coelom lies within the abdominal blood

REYNOLDS: SCAPHOPOD HEART 139

Fig. 1. Longitudinal section through the perianal sinus (pa), kidney (n), and anterior portion of stomach (s) and pericardium (arrowheads) (ab,
abdominal sinus; mc, mantle cavity; pc, pericardial cavity; r, rectum). Scale bar = 0.1 mm. Fig. 2. Oblique cross section of the perianal sinus
(pa), showing traversing muscular trabeculae (arrowheads) (mc, mantle cavity; r, rectum). Scale bar = 40 um. Fig. 3. Muscle cells of the perianal
sinus (Sm) and the pericardium (pm). Note neural process adjacent to perianal sinus musculature (arrowhead) (h, haemocoel; pc, pericardial
cavity; pe, pericardial epithelial cell). Scale bar = 1 um. Fig. 4. Cytoplasmic extensions of a pericardial epithelial cell overlying a muscle cell
of the perianal sinus (arrowheads, dense bodies; arrow, attachment plaque; g, glycogen granules; h, haemocoel; jsr, junctional sarcoplasmic

reticulum; m, mitochondrion; pc, pericardial cavity; sr, sarcoplasmic reticulum; th, thick myofilaments). Scale bar = 0.5 um. Fig. 5. Thick
myofilaments of the smooth perianal sinus muscle cell. Note axial periodicity within the filaments. Scale bar = 0.2 um.

sinus, ventral to the stomach, and extends from the posterior
end of the stomach to the kidneys and perianal sinus, to which
it adheres anteriorly (Figs. 1, 6-8). The ventral pericardial
epithelium is always in close contact with the body wall, while
irregular infoldings of the dorsal pericardial wall project into
the coelomic cavity. There is no myocardium or any type of
endothelium within these foldings (Figs. 1, 7, 8); the only
musculature adjacent to the pericardium is that of the body
wall ventrally and perianal sinus anteriorly (Figs. 6, 7). A con-
nection exists between the pericardial coelom and the right
kidney (Fig. 9), although it was not found in all specimens.

The pericardial wall is composed of three cell types:
simple flat epithelium, interspersed with muscle cells, and
modified in the region adjacent to the perianal sinus to in-
clude podocytes. The arrangement and ultrastructure of
epithelial and muscle cells is similar throughout the peri-
cardium (Fig. 6). The epithelial cells (Fig. 10) typically possess
a cell body with a small amount of cytoplasmic material sur-
rounding the nucleus. The cell bodies extend into the pericar-
dial cavity, with the continuous basal lamina lining the
haemocoel. Thin cytoplasmic branches extend between cell
bodies and contain one or a few isolated mitochondria in ad-
dition to a- (15 nm) and £- (37 nm) glycogen granules (Figs.

10-12). Desmosomes, with an intercellular distance of 9-15 nm,
occur frequently where epithelial cell junctions appose the
basal lamina (Fig. 11) but were not observed in areas where
cytoplasmic extensions overlap adjoining muscle cells (Fig.
12). Adjoining plasmalemmae are not highly infolded and do
not interdigitate (Fig. 12).

The pericardial musculature is arranged as trabeculae
which run in an entirely transverse direction, and are discon-
tinuous in both anterior-posterior and lateral axes of the
pericardium. The width of the trabeculae varies between 1
and 10 um (Figs. 13, 14). Muscle cells are interspersed be-
tween the epithelial cells and typically underlie extensions
of the epithelial cytoplasm (Figs. 12, 15-18). Adjoining plas-
malemmae of the two cell types have an intercellular space
of 7-15 nm within which no extracellular material has been
observed (Figs. 12, 15, 16, 18). Desmosomes occur at cell junc-
tions apposing either the basal lamina or coelomic space, and
have an intercellular width of 9-15 nm (Fig. 16, 18). Both cell
types are separated from the haemocoel by a continuous
basal lamina, 18-40 nm thick (Figs. 12, 15-18). A thin layer
of collagen fibrils, varying in thickness from 0.11-0.53 um, is
often associated with the basal lamina (Fig. 15). Neural
elements are found adjacent to the muscle cells (Fig. 17).

140 AMER. MALAC. BULL. 7(2) (1990)

Fig. 6. Schematic diagram showing the relative positions of the
perianal sinus (pas), pericardium (pc) and kidneys (n) (ab, abdominal
sinus; bw, body wall; dd, digestive diverticulum; pd, podocytes; pe,
epithelial cell of the pericardium; pm, muscle cell of the pericardium;
r, rectum; A-B indicates cross-sectional view represented in figure 7).

Fig. 7. Schematic diagram showing the relative positions of the
stomach (s), pericardium (pc) and mantle cavity (mc) (ab, abdominal
sinus; bw, body wall; dd, digestive diverticulum; ma, mantle; pe,
epithelial cell of the pericardium; pm, muscle cell of the pericardium;
rm, retractor muscle; C-D indicates frontal section view represented
in figure 6) (See also figure 17).

Muscle fibres of the pericardium fixed in a contracted
state show near-alignment of dense bodies (0.12-0.14 um
length, 46-58 nm width) into Z-lines, with the intervening thick
myofilaments creating A and | lines in a loose sarcomeral
structure, approximately 1 wm in length (Fig. 19). Occasional
attachment plaques were observed anchoring the filaments to
the sarcolemma (Fig. 19). The muscle cells have a diameter
at the nucleus of 3-35 um (Fig. 20). Thick and thin
myofilaments do not appear to have a regular arrangement
with respect to each other, and have diameters of 18-33 nm
and 6-7.5 nm respectively (Figs. 16, 18). Profiles of rough and
smooth sarcoplasmic reticulum are present as are those of
junctional sarcoplasmic reticulum (Figs. 16, 18, 19). A small

quantity of a- and B- glycogen granules is present within the
cytoplasm of the cell periphery (Fig. 16). Clusters of mito-
chondria are positioned between the contractile elements and
sarcolemma; sarcolemmal width is 7-15 nm (Fig. 21).

The pericardium contracts independently of the
perianal sinus in a regular though discontinuous manner;
there is neither a gradual peristalsis of the pericardium nor
a simultaneous uniform contraction of all muscle fibres. The
contractions occur in an anterior to posterior progression of
2-3 discrete constrictions of the dorsal pericardial wall, the
wholly transverse orientation of the muscle fibres producing
a single localized transverse constriction which is released
as another, posterior to this, is produced. The posterior end
of the pericardium appears to lie free within the abdominal
sinus and moves in an anterior-posterior direction due to con-
traction of the muscle fibres. The anterior wall remains in
close contact with the kidneys and perianal sinus, as the ven-
tral pericardial wall does with the body wall. Very infrequent-
ly a slight contraction of the stomach was observed.

The third cell type of the pericardium is the podocyte,
which is characterized by the presence of pedicels and
fenestrations in the cytoplasmic branches of the pericardial
epithelium (Figs. 22-26). The cell type is not widespread and
has only been observed in areas apposed by smooth
musculature in the region of the perianal sinus, i.e. the antero-
ventral portion of the pericardium (Figs. 6, 23, 24). Fenestra-
tions, 13-32 nm in width, are distributed along cytoplasmic
extensions (Figs. 22-26), and raised pedicels have also been
observed (Fig. 25). In some sections, diaphragms in the form
of electron opaque strands bridge the fenestration (Fig. 26).
In all cases, the fenestrations overlie the basal lamina; there
is no evidence of an apposing collagen layer. No micro-
villi line the luminal surface of these or any cells of the
pericardium.

DISCUSSION

STRUCTURE OF THE MOLLUSCAN HEART

The anatomy of the molluscan heart generally consists
of a single ventricle and one or two auricles which usually
correspond to the number of ctenidia. The whole is enclosed
within a pericardium, and the ventricle in the Bivalvia and
some Gastropoda is traversed by the rectum (Jones, 1983).
At the height of molluscan heart organization, the Ceph-
alopoda possess a complex systemic heart associated with
a closed circulatory system (Wells, 1983); at the other extreme,
the Scaphopoda have been described as having a rudi-
mentary heart, relying on body musculature for circulation
through a series of sinuses (Hill and Welsh, 1966). The
molluscan heart lies freely within the pericardium, although
the pericardial cavity does not extend dorsally over the ven-
tricle in the Neomeniomorpha (Salvini-Plawen, 1985) and
the bivalve Pteria (White, 1942). In some bivalve species the
dorsal wall of the ventricle remains attached to the pericar-
dium by connective tissue (Narain, 1976). The presence of
the pericardium is critical to circulatory function; the pressure
of the pericardial fluid is normally less than that of the blood

REYNOLDS: SCAPHOPOD HEART 141

Fig. 8. Longitudinal section through the pericardium (pc), stomach (s), perianal sinus (pa) and kidney (n) (arrowheads, anterior and dorsal
pericardial walls; ab, abdominal sinus; dd, digestive diverticulum; i, intestine; mc, mantle cavity). Scale bar = 0.15 mm. Fig. 9. Longitudinal
section showing connection between the pericardial cavity (pc) and the right kidney (n) (dd, digestive diverticulum; mc, mantle cavity; s, stomach).
Scale bar = 50 um. Fig. 10. Pericardial epithelial cell (arrow, basal lamina; h, haemocoel; m, mitochondrion; n, kidney; pc, pericardial cavity).
Scale bar = 2.5 um. Fig. 11. Cytoplasmic extensions of the pericardial epithelium (arrowhead, desmosome; h, haemocoel; pc, pericardial
cavity). Scale bar = 0.3 um. Fig. 12. Epithelial (pe) and muscle cells (pm) of the pericardium (arrowhead, basal lamina; arrow, myofilaments;
h, haemocoel; pc, pericardial cavity; sr, sarcoplasmic reticulum). Scale bar = 0.6 um.

in the heart, and cardiac refilling is maintained by a volume-
compensating mechanism as originally proposed by Ramsay
(1952) and Krijgsman and Divaris (1955), and reviewed by
Jones (1983). The pericardium is drained by one or two
renopericardial canals, which connect the lumina of the
pericardium with those of the kidneys (Martin, 1983).

At the cellular level, the molluscan heart consists of
an epicardium, which rests on a basal lamina, and an inner
loose myocardium; an endothelium is lacking (Narain, 1976;
@Okland, 1980; Jones, 1983) except in cephalopods, where it
is incomplete (Jensen and Tjonneland, 1977). While in some
cases the epicardial cells possess microvilli and can also have
a secretory role (Kling and Schipp, 1987), the ventricular
epicardium generally consists of a continuous, simple
epithelium. Podocytes are usually concentrated within the
auricular epicardium, and are characterized by the presence
of numerous thin cytoplasmic extensions termed pedicels,
which are aligned in a parallel array over the continuous basal
lamina, between the haemocoel and coelomic spaces
(Andrews, 1976; Pirie and George, 1979; Okland, 1980). The
gaps between the pedicels create fenestrations in the
epithelium or ultrafiltration slits; the basal lamina separates
the haemocoel from pericardial coelom, and acts as the func-
tional filter (Andrews, 1981; Morse, 1987). In many bivalves,

gastropods and cephalopods the ultrafiltration slits are
bridged by slit diaphragms (Boer and Sminia, 1976; Andrews,
1979; Schipp and Hevert, 1981; Meyhdfer et a/., 1985),
although these have not been found in the podocytes of
chitons (Okland, 1980). Portions of the auricular epicardium
or pericardial epithelium are elaborated in many species, par-
ticularly in bivalves, to form pericardial glands (White, 1942),
within which extensive areas of podocytes have been found
(Meyhofer et al., 1985). A similar development of the
haemocoel-pericardial interface is seen in the branchial heart
appendages of coleoids and the pericardial appendages of
nautiloid cephalopods (Fiedler and Schipp, 1987). The
molluscan pericardium consists primarily of simple epithelial
cells, although there have been few detailed ultrastructural
studies. The polyplacophoran pericardial epithelium is con-
tinuous with, while differing from, the ventricular and auricular
epicardium (Okland, 1980); it also possesses muscle cells and
is pulsatile (Greenberg, 1962; Okland, 1981).

INTERPRETATIONS OF SCAPHOPOD CIRCULATORY
STRUCTURES TO DATE

Two structures have been proposed as representing
the scaphopod heart: (i) the perianal sinus, which has been
described as having morphological similarities to (Lacaze-

142 AMER. MALAC. BULL. 7(2) (1990)

Fig. 13. Dorsal pericardial wall, viewed from the pericardial cavity. Anterior is to the top of the photomicrograph. Note lateral orientation of
muscle fibres. Scale bar=0.15 mm. Fig. 14. Dorsal pericardial wall, viewed from the pericardial cavity. Anterior is to the top of the photomicrograph.
Note the discontinuity of the pericardial muscle cells (pm) along anterior-posterior and lateral axes. Scale bar = 40 um.

Fig. 15. Epithelial (pe) and muscle cells (pm) of the pericardium (arrowhead, basal lamina; arrow, collagen fibres; h, haemocoel; pc, pericardial
cavity; sc, subsarcolemmal cisternae). Scale bar = 2 um. Fig. 16. Junction of epithelial (pe) and muscle cells (pm) of the pericardium (ar-
rowheads, basal lamina; arrow, desmosome; h, haemocoel; sr, sarcoplasmic reticulum; th, thick myofilaments). Scale bar = 0.5 um. Fig. 17.
Longitudinal section of dorsal (top) and ventral (bottom) pericardial walls and body wall (bw) (*, muscle cells of the pericardium; h, haemocoel;
mc, mantle cavity; ne, nerve process; pc, pericardial cavity; sc, subsSarcolemmal cisternae). Scale bar = 10 um. Fig. 18. Junction of epithelial
and muscle cells of the pericardium (arrowheads, basal lamina; arrow, desmosome; h, haemocoel; jsr, junctional sarcoplasmic reticulum; ne,
nerve process; pc, pericardial cavity; sr, sarcoplasmic reticulum; th, thick myofilaments). Scale bar = 0.7 um. Fig. 19. Longitudinal section
through pericardial muscle cell. Note loose sacromeral structure formed by alignment of dense bodies. Scale bar = 0.15 um. Fig. 20. Oblique
cross section through pericardial muscle cell (arrowhead, basal lamina; h, haemocoel; nu, nucleus; pc, pericardial cavity). Scale bar = 3 um.

REYNOLDS: SCAPHOPOD HEART 143

Duthiers, 1857) and homology with (Fol, 1889) the bivalve ven-
tricle; and (ii) the dorsal infolding of the pericardium, ventral
to the stomach, described by Plate (1891, 1892), Boissevain
(1904), and Distaso (1905). Lacaze-Duthiers (1857) did not
discuss the relationship between the perianal sinus and
molluscan heart in any detail. Fol (1889), however, based
their homology on structural similarities, describing the posi-
tion of the sinus in relation to the rectum, the musculature
and rhythmic contractions of the sinus, and an endothelium
which Plate (1892) and Boissevain (1904) later refuted.
Lacaze-Duthiers (1857) placed the major responsibility for
movement of the blood with the large pedal sinus, while Fol
(1889) agreed that the perianal sinus makes little contribu-
tion to circulation.

The studies by Plate (1891, 1892), which were con-
firmed by Boissevain (1904) and Distaso (1905), described a
contractile vessel ventral to the stomach, surrounded by the

pericardial coelom. Plate (1892) stated that the heart is ex-
traordinarily simple, with no chambers or vessels, and lacks
a strong development of musculature. He found that the
pericardial and heart walls did not differ histologically, and
were composed of epithelial cells with very thin, parallel and
regularly arranged fibres. While Plate (1892) suggested that
these could be muscle fibres, he concluded that this struc-
ture could not act as a center of propulsion for the circulatory
system.

CIRCULATORY STRUCTURES IN
DENTALIUM RECTIUS

In Dentalium rectius, there is no evidence of the
ultrastructural features associated with the typical molluscan
heart (i.e. a myocardium with an associated epicardium) in
a position ventral to the stomach and enclosed by the peri-

Fig. 21. Junction of pericardial muscle cells (m, mitochondrion; pc, pericardial cavity; th, thick myofilaments). Scale bar = 1 um. Fig. 22.
Podocytes in the pericardium. Note fenestrations in the pericardial epithelium apposed by basal lamina (arrowheads) (nu, nucleus; pc, pericar-
dial cavity). Scale bar = 1 wm. Fig. 23. View of perianal sinus muscle cells (sm) and pericardium. Note the highly infolded cytoplasmic exten-
sions of the pericardium overlying the perianal sinus musculature, and the fenestrations apposed by basal lamina (arrowheads) (h, haemocoel:;
pc, pericardial cavity). Scale bar = 1 wm. Fig. 24. Muscle cell of perianal sinus (sm) and fenestrations (arrowheads) in the overlying pericar-
dium (g, glycogen granules; th, thick myofilaments). Scale bar = 0.7 um.

144 AMER. MALAC. BULL. 7(2) (1990)

cardium as described by Plate (1891, 1892), Boissevain (1904)
and Distaso (1905) for other species of Dentalium. Without
the benefit of ultrastructural study, it is likely that these early
investigators considered the contractile dorsal pericardial wall
as a ventricular epicardium. The lack of a myocardium or any
ultrastructural differentiation of this portion of the pericardial
epithelium discounts this interpretation, and no evidence sup-
porting homology with the molluscan ventricle exists. While
the scaphopod stomach is described as possessing a
muscular tunic (Salvini-Plawen, 1988), the musculature in D.
rectius is discontinuous and closely applied to the stomach
wall and as such is not associated with the pericardium and
does not enclose a portion of the haemocoel.

The homology of the pericardial coelom described by
Lacaze-Duthiers (1857), Plate (1891, 1892), Boissevain (1904)
and Distaso (1905) with that of other molluscs is based sole-
ly on its general anatomy, i.e. a closed sac composed of
squamous epithelium within the haemocoel. Ultrastructural
features of the pericardial epithelium in Dentalium rectius sup-
port this homology; the presence of long cytoplasmic
branches, desmosomes and few other organelles suggest a
simple delimiting epithelium, similar to that found in other
molluscan peri- and epicardia. Furthermore, an excretory role
inferred from the presence of podocytes and a renopericar-
dial connection also supports this homology. On this basis
the term pericardium should be retained in describing this
structure in scaphopods.

The arrangement of musculature in the Dentalium rec-
tius pericardium suggests transverse contractions of the dorsal
pericardial wall, which are supported by the observations of
live animals. The contractile pericardia in the Polyplacophora
have been suggested by Okland (1981) to function in the cir-
culation of pericardial fluid as part of the excretory system,
with little effect on the contraction mechanisms of the heart.
In Tonicella, the epithelial and muscle cells are separated by
collagen and basal lamina, which is, however, continuous with
the basal lamina lining the epithelial cells (Okland, 1981). In
comparison, no extracellular material exists between cell types
in the pericardium of D. rectius, and the basal lamina is limited
to an unbranching layer between the pericardial elements and
haemocoel. Contractions of the dorsal pericardial wall in D.
rectius undoubtedly contribute to the circulation of blood
through the relatively large abdominal sinus. These contrac-
tions do not lead directly to ultrafiltration of blood through the
dorsal pericardial wall due to the absence of podocytes in this
area of the pericardium. However, it is possible that local in-
creases in blood pressure could be transferred anteriorly to
the perianal sinus and ultimately facilitate ultrafiltration
via the podocytes lining the perianal sinus. The irregular in-
vaginations of the dorsal pericardial wall of D. rectius seen
in section and considered by Plate (1891, 1892), Boissevain
(1904) and Distaso (1905) in other Dentalium species to be
the heart, are due to the state of contraction of the dorsal
pericardial wall at fixation and do not represent a permanently
enclosed contractile vessel. The ventral pericardial wall was
always observed in close adherence to the body wall in both
fixed and live material.

The presence of podocytes is a development of the

pericardial epithelium that is commonly observed and inter-
preted in other molluscan classes as the site of ultrafiltration
of blood and production of primary urine. While ultrastructural
features such as apposition of basal lamina, fenestration width
and the presence of slit diaphragms in the podocytes of
Dentalium rectius are consistent with such a function, there
is no evidence of an extensive array of pedicels and ultrafiltra-
tion slits as seen in representatives of some other molluscan
classes (Andrews, 1979; MeyhOfer et a/., 1985). Thus, the area
available for ultrafiltration appears to be quite limited in D.
rectius.

The renopericardial connection with the right kidney
is small (20 »m in diameter), and was only noted in inciden-
tal thin (1m) sections. A left renopericardial canal in
Dentalium was described by Distaso (1905), although it ap-
pears from reading his account that the dorso-ventral orien-
tation of the animal is opposite to that generally accepted by
other investigators of the Scaphopoda. Therefore, consider-
ing the mantle cavity as ventral and the larger aperture as

Fig. 25. Podocytes of the pericardium. Note raised pedicels (arrows)
(h, haemocoel; pc, pericardial cavity). Scale bar = 0.3 um. Fig. 26.
Podocytes of the pericardium. Note fenestrations in epithelium ap-
posed by basal lamina (arrowheads) and bridged by diaphragms (ar-
rows) (h, haemocoel; pc, pericardial cavity). Scale bar = 0.3 um.

REYNOLDS: SCAPHOPOD HEART 145

anterior, Distaso (1905) had, in fact, also described a connec-
tion between the pericardium and right kidney.

The ultrastructure of the perianal sinus in Dentalium
rectius does not differ significantly from that of smooth
molluscan cardiac muscle. Thick myofilaments have an ax-
ial periodicity resembling paramyosin, while myofibre size, the
arrangement of glycogen and mitochondria, and the develop-
ment of the sarcoplasmic reticulum is similar to that found
in the cardiac musculature of the bivalves Venus (Kelly and
Hayes, 1969), Elliptio (Rutherford, 1972) and Geukensia (Watts
et al., 1981), and of the polyplacophorans Lepidopleurus and
Tonicella (Okland, 1980). This is in contrast with the oblique-
ly striated heart musculature of gastropods and cephalopods,
as reviewed by Kling and Schipp (1987). The position of the
sinus in D. rectius in relation to the rectum parallels that
found in the bivalve ventricle. Also, the presence of travers-
ing muscular trabeculae, which produce the regular contrac-
tions of the sinus (this study; Fol, 1889; Plate, 1892; Fischer-
Piette and Franc, 1968), is also found in the ventricles of many
bivalves and gastropods (Narain, 1976; Okland, 1982; Jones,
1983).

Whether the perianal sinus, pericardium and kidneys
in Dentalium rectius have an associated ontogenesis reflec-
ting the developmental pattern of the heart, pericardium and
kidneys from a common anlagen as seen in other classes of
molluscs (Raven, 1966; Moor, 1983) with a subsequent
movement of the pericardium from a position surrounding the
ventricle to one more posterior to it, awaits further informa-
tion on scaphopod organogenesis. If an homology between
these organs exists, the lack of aortae, auricles or valves of
any description, and the limited apposition of the pericardium,
indicate a much reduced heart compared to that found in other
molluscan classes. This reduction could have developed as
a consequence of altered circulatory requirements, due in
large part to the loss of ctenidia from the uniquely modified
scaphopod mantle cavity.

Unidirectional flow of blood through the perianal sinus
is not maintained (pers. obs.). While contractions could pro-
duce local pressures capable of driving limited ultrafiltration,
it is unlikely to be capable of overcoming peripheral resistance
of the circulatory system, or of even contributing significant-
ly to circulation of the blood. As a consequence of the con-
tractions of the foot and body musculature, however, the
perianal sinus may serve a role in facilitating equilibration of
pressure gradients between the pedal and abdominal blood
sinuses.

In conclusion, there is no evidence for a heart within
the pericardium of Dentalium as interpreted by Plate (1891,
1892), Boissevain (1904), and Distaso (1905). Ultrastructural
features in Dentalium rectius suggest that there could be an
homology of the perianal sinus and pericardium with the heart
and pericardium of other molluscs. Studies of scaphopod
organogensis are necessary to confirm this. The contractility
of the dorsal pericardial wall and the perianal sinus may
facilitate ultrafiltration of the blood via podocytes, which are
limited to the anterior portion of the pericardium and overlie
the perianal sinus. Further study into the blood pressures
created by these and other contractile structures of the

Dentalium haemocoel is necessary to further delineate their
respective contributions to circulation.

ACKNOWLEDGMENTS

| am grateful to D. McHugh and R. Marx for assistance with
translation of the German texts. | thank Dr. A. R. Fontaine and D.
McHugh for critically reading earlier drafts of the manuscript, and
two anonymous reviewers for useful comments. This research was
funded in part by a University of Victoria Fellowship.

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Date of manuscript acceptance: 12 October 1989.

DIET AND HABITAT UTILIZATION IN A NORTHEASTERN
PACIFIC OCEAN SCAPHOPOD ASSEMBLAGE

RONALD L. SHIMEK'
BAMFIELD MARINE STATION
BAMFIELD, BRITISH COLUMBIA
CANADA VOR 1BO

ABSTRACT

The diets of Dentalium rectius Carpenter, 1864, Pulsellum salishorum Marshall, 1980, and Cadulus
aberrans Whiteaves, 1887 were determined by examination of buccal pouch contents of specimens
collected and examined quarterly from December 1983 to December 1984. D. rectius was omnivorous,
ingesting a wide variety of food items including sediment particles, fecal pellets, kinorhynchs, and various
invertebrate eggs; however, foraminiferans were the most numerous prey. D. rectius was most abun-
dant in a silty area containing about 10% organic material by weight; however, it was found in all areas
sampled. D. rectius ranged in abundance from about 5 animals/m2 in clean sand to about 66 animals/m2
in silt. Foraminiferans were rare where D. rectius was most abundant.

Cadulus aberrans preyed preferentially upon the foraminiferans Cribrononion lene and Rosalina
columbiensis. The robust foraminiferan E/phidiella hannai was readily accepted as prey, while the fragile
Florilus basispinatus were taken less frequently than expected, as were Buliminella spp. C. aberrans
was found most frequently in sandy substratum consisting of about 5% organic material by weight.
Foraminiferans were common in this habitat. The average density of C. aberrans was about 10
animals/m2.

Pulsellum salishorum was a dietary specialist preying on the foraminiferan Cribrononion lene.
P. salishorum was relatively uncommon (6 animals/m2) but evenly distributed in all habitats examined.

Due to selective predation on foraminiferans, scaphopods appear to alter relative abundances
and size-frequency distributions of their prey populations. The numerical dominance of Florilus
basispinatus, Buliminella elegans, and Buliminella exilis and the relative rarity of Cribrononion lene are
probably direct results of scaphopod predation. Most of the prey items were less than 300 xm in diameter.

Dentalium rectius is able to thrive in areas of low populations of foraminiferans by utilizing alter-
native foods. Predation by D. rectius and Pulsellum salishorum in these habitats probably causes these

low populations.

Scaphopods, uncommon members of most shallow-
water marine ecosystems, have seldom been studied. The
diets and some natural history attributes are known for a few
species, usually based on small sample sizes or limited to
short periods of observation (Davis, 1968; Gainey, 1972; Mc-
Fadien, 1973; Bilyard, 1974; Poon, 1987). Generally, previous
studies were made on members of the order Dentalioida, with
few observations on species in the other scaphopod order,
Gadilida (Davis, 1986; Carter, 1983; Poon, 1987).

Scaphopods are the only wholly infaunal molluscan
class and are relatively abundant in deep-sea communities.
They are also abundant in the unconsolidated sediments of
the fjord systems north of the Strait of Juan de Fuca on the

1Current address: 25022 144th Place S.E., Monroe, Washington
98272, U.S.A.

West Coast of North America (Shimek, 1988, 1989). In these
areas, with the exception of minute bivalves such as Axinop-
sida serricata (Carpenter, 1864), scaphopods are often the
dominant mollusks with several sympatric species.

Morton (1959) referred to the Scaphopoda as the ‘‘most
uniform group” of mollusks. Scaphopods are generally con-
sidered to be predators upon foraminiferans (Lacaze-Duthiers,
1856, 1857; Morton, 1959; Dinamani, 1964; Fisher-Piette and
Franc, 1968; Gainey, 1972; Bilyard, 1974; Taib, 1980; Carter,
1983; Poon, 1987). | examined several scaphopod com-
munities in Barkley Sound on the southwest side of Vancouver
Island, British Columbia to determine diet and habitat utiliza-
tion in order to address the possibility of competition for either
food or habitat.

| tested the hypothesis that these communities con-
tained representatives of different species that were eating

American Malacological Bulletin, Vol. 7(2) (1990):147-169

147

148 AMER. MALAC. BULL. 7(2) (1990)

similar prey and living in the same habitats. To exan ‘ne this
general hypothesis, | asked several specific gestions about
diet. Do these animals eat the same general category of prey?
Within those categories, do these animals prey upon
representatives of the same species? Are the prey similar in
size or shape? Does the diet vary seasonally or with reproduc-
tive condition of the predator? Do the prey vary with the habitat
or are scaphopods feeding on similar prey among various
habitats? | asked similar questions about the habitats where
the scaphopods were collected. What were the physical
characteristics of the habitats? Could the distribution of either
the scaphopods or their prey be correlated with any particular
physical parameter of the habitat?

MATERIALS AND METHODS

Scaphopods for this study were collected at three sites
in Barkley Sound (Table 1). The most abundant scaphopods
were Dentalium rectius Carpenter, 1864, Cadulus aberrans
Whiteaves, 1887, and Pulsellum salishorum Marshall, 1980.
Other sympatric species were C. californicus Pilsbry and
Sharp, 1898, C. to/miei Dall, 1897, and D. pretiosum Sowerby,
1860. Some of the latter three species were more common
in other habitats, but these were not included in this study.

Quantitative collections were made using a 0.1m2
Petersen bottom grab. Two replicate samples were collected
from each station in December, 1983 and March, June,
September and December, 1984. After every haul, the grab
was inspected to insure adequate substratum penetration and
complete jaw closure. The grab was further examined to check
for evidence of sample loss due to winnowing. If the grab func-
tioned incorrectly, that sample was discarded and a replace-
ment taken. Each sample was deposited in a sorting tray, and
a il subsample of substratum was retained for later particle
size and composition analyses. The remaining sediment was
wasned gently through a 0.5 mm screen and all scaphopods
were retained.

At each site a beam trawl or anchor dredge was used
to collect additional scaphopods. Specimens to be examined
alive were cleaned of any adherent sediment, placed in clean
sea waiter, and returned to Bamfield Marine Station. During
transport and handling, the animals were maintained below
15°C. Exposure to higher temperatures is generally lethal to
scaphopods (Shimek, 1988). At the laboratory specimens were
maintained in separate chambers in water tables.

Specimens not treated as above were fixed in 10% buf-
fered formalin immediately after collection. After 24 to 48 hours
they were rinsed with fresh water and transferred to 70% EtOH
for storage. If buccal contents were to be examined, 1.0%
Rose Bengal, by weight, was added to the alchohol in order to
facilitate identification of organic materials (Bilyard, 1974;
Shimek, 1988).

Scaphopod shells were measured (Shimek, 1989) and
the soft parts removed for analysis of buccal contents as
described by Bilyard (1974). All buccal pouch contents were
enumerated and measured. Measurements taken varied with
shape of the food items. The measurement was typically made
along the semimajor axis, i.e. the second longest measure-

ment. (During feeding the longest dimension of the prey, the
major axis, is oriented normal to the plane of the buccal open-
ing, thus the maximum distention of the buccal opening has
to accommodate the second longest dimension of the prey.)
Prey items were identified using available works (Cockbain,
1963, Lankford and Phegler, 1973; Gallagher, 1979; Kozloff,
1987).

Buccal pouch clearance rates were determined by
periodic observation of living specimens. Immediately upon
return to the laboratory, live scaphopods were put onto clean
substratum from their native habitat. This sediment was
previously treated with fresh water for several days in order
to kill resident infauna. The sediment was then placed in
miniature aquaria held inside of larger aquaria within a flow-
through sea-water system. Perforations in the bottoms of the
the miniature aquaria were coverd with 63 »m mesh plastic
screen. The tops of the smaller aquaria were held above the
water level in the water table and running sea water was sup-
plied to them.

Five specimens of a given species of scaphopod were
put into each aquarium; only those that burrowed complete-
ly into the substratum were used to determine buccal
clearance rates. Starting at 24 hours after collection, some
species were removed from the substratum and fixed for later
examination. Additional specimens were fixed at 6 hr inter-
vals thereafter. All scaphopods in a given aquarium were
removed simultaneously. The buccal pouch contents were ex-
amined as indicated above.

Substratum was analyzed for particle size distribution
following the methods of Holme and Mcintyre (1971). A
modified wet-sieve method was used to determine particle
sizes to 0.63 «m. The total silt-clay fraction was determined
by evaporation and weighing, but was not partitioned further.

The sediment organic content was estimated by deter-
mination of total volatile solids. Approximately 25 g of sedi-
ment was dried in a tared, pre-oxidized, aluminum pan and
total weight determined. The pan and sediment were then
heated in a muffle furnace at 500°C for 24 hr. The sediment
and pan were allowed to cool in a desiccator and re-weighed.
The difference between the two weights was used as weight
of the total volatile solids.

Potential prey items were isolated from paired replicate
substratum subsamples taken from the 11 sample of quan-
titatively collected sediment. After sediment for particle size
analysis was removed, the remaining material was
homogenized by stirring with a small amount of added sea
water. A small aliquot (40 to 90 ml) of the homogenate was
removed and fixed in 10% sea-water buffered formalin with
rose bengal added. After 24 hours, the samples were wash-
ed through a 63 »m mesh screen and stored in 70% ethanol
until examination.

Sediment samples were examined at 10 to 40
diameters using a Wild M-5 stereomicroscope and all infauna
were enumerated and measured. Foraminiferan tests were
seen commonly, but were not identified, measured, or
enumerated. Similarly, time constraints did not allow the
numerous other fragmentary remains, e.g. molluscan shells,
scaphopod shells, heart urchin spines, and polychaete setae,

SHIMEK: SCAPHOPOD DIETS

to be measured, even though these items contribute to the
gut contents of some scaphopods. The potential food value
of these items is unknown. Fine inorganic particles and fecal
pellets, the most common non-animal constituents of the sedi-
ment, were not enumerated even though they were found in
some buccal contents. The relative proportion of each taxon
in the infauna or the buccal contents was calculated by
dividing the number of individuals of a particular taxon by the
total number of individuals in that sample.

For statistical comparisons, potential prey from a par-
ticular habitat were defined as all individuals of all taxa that
had been found at least once as part of the dietary intake of
a scaphopod. Thus, all foraminiferans were considered to be
potential prey, as were all bivalves; polychaetes and
nematodes were not.

Most statistical analyses were performed using
STATGRAPHICS (STSC, 1986, 1987, 1988). For analyses of
variance, proportions were transformed by arcsine square-
root transformation to decouple the variance from the means
(Sokal and Rohlf, 1981). [Proportions approximate binomial
distributions and the variance is a function of the mean which
introduces bias into the analysis of variance (ANOVA). The
arcsine square-root transformation prevents this.] The
Shannon-Wiener diversity index (H) and the evenness index
(J) were calculated where applicable (Poole, 1974).

RESULTS

Stations sampled differed in depth (Table 1), particle
size distribution and volatile solids content (Figs. 1, 2, Table

149

2). Particle size distribution and total volatile solids differed
significantly among stations, although not seasonally within
a single station (Table 2).

The number of quantitative grabs varied among sta-
tions. Nonetheless, sufficient samples were taken to adequate-
ly assess scaphopod abundances, which varied among and
within stations by species (Table 3). More scaphopods were
found per unit area at the Sandford Island station than at the
other two sites. Here Dentalium rectius was numerically domi-

Table 1. Scaphopod study sites and collection areas in Barkley Sound,
Vancouver Island, British Columbia.

A. Station M - Mayne Bay, Northeastern Corner of Barkley Sound.
Corners of the sample collection area: 48° 58.8’ N, 125° 18.9’ W;
48° 59.0’ N, 125° 19.2’ W; 48° 586’ N, 125° 20.1’ W;
48° 58.4’ N, 125° 19.7’ W.
Centroid of the sample collection area: 48° 58.7’ N, 125° 19.5’ W.
Depth range of the sample collection area: 35-40 m.

B. Station S - Off Sandford Island, in Imperial Eagle Channel.
Corners of the sample collection area: 48° 52.3’ N, 125° 11.4’ W,
48° 52.5’ N, 125° 11.7’ W, 48° 53.1’ N, 125° 11.1’ W,
48° 52.8’ N, 125° 11.2’ W.
Centroid of the sample collection area: 48° 52.7’ N, 125° 11.4’ W.
Depth range of the sample collection area: 75-80 m.

C. Station T - Trevor Channel, near Diana Island. Corners of the
sample collection area: 48° 50.0’ N, 125° 10.8’ W, 48° 50.2’ N,
125° 10.0’ W, 48° 49.3’ N, 125° 11.7’ W, 48° 49.1’ N, 125° 11.5’ W.
Centroid of the sample collection area: 48° 49.7’ N, 125° 11.0’ W.
Depth range of the sample collection area: 30-110 m.

Table 2. Tests of significance of the differences in station sediment parameters.

Sum of
Squares

Source of Variation

df.

A. Analysis of variance for proportional sediment weights (proportions are arcsine-square root transformed).

Main Effects 13.931
Particle Size 13.794
Station 0.104
Month 0.032

2-Factor Interactions 2.786
Particle Size x Station 2.519
Particle Size x Month 0.253
Station x Month 0.014

Residual 1.226

Total 17.942

13
8
2
3

46
16
24
6
156
215

Mean F-ratio P
Square

1.072 136.352 <0.0001
1.724 219.396 < 0.0001
0.052 6.645 0.0017
0.011 1.372 0.2533
0.061 7.705 < 0.0001
0.157 20.030 <0.0001
0.011 1.341 0.1463
0.002 0.296 0.9382
0.008

B. Analysis of variance for proportional total volatile solids (proportions are arcsine-square foot transformed).

Main Effects 2.966
Grain Size 2.753
Station 0.175
Month 0.038

2-Factor Interactions 3.936
Grain Size x Station 1.663
Grain Size x Month 1.759
Station x Month 0.514

Residual 4.166

Total 11.068

13

0.228 8.544 < 0.0001
0.344 12.886 < 0.0001
0.087 3.272 0.0405
0.013 0.478 0.6981
0.086 3.204 < 0.0001
0.104 3.891 <0.0001
0.073 2.745 0.0001
0.086 3.208 0.0053
0.027

150 AMER. MALAC. BULL. 7(2) (1990)

[Slo lonta| ok Tahal [rE Se le (i) (aa i Pea i es en
ab
[
@.8+ + MAYNE BAY al
= <> SANDFORD ID.
4 - TREVOR CH.
k-
&
a 0.65 5 4
v4 )
a va
4 Z
z /
q e:4 / ar
ae
= / |
=) [ y 1
oO 4 {
| 4
6.2 | 4
of /
ea ct oy Shor
Oo yf - Sl ia
Pelli ct mens ee ee
! tof oj ij | j riitli SS) CS DS ee ee es jeve este eae TT 4 i onl ees J
=3 -2 -1 7) 1 2 3 4
PHI
Fig. 1. Sediment particle size frequency distribution for all habitats,
means + 1 standard deviation indicated [Phi = -logz (sediment par-
ticle diameter)].
6.5)
+ MAYNE BAY
8.4) < SANDFORD ID.+
-} TREVOR CH. |
| |
1
= | |
5 9-3 |
H i be
=
na |
0 | |
o | | |
O
© y 1
o 0.2 t owes
{Nt a 1
4 ys i if |
LW |
i I/ \ ce
@.1 t e Bo
| Ki I
| Ba |
)
= = ej 1 | Jt re er | i
re] -2 =A ) 1 2 3 4
PHI

Fig. 2. Sediment total volatile solids by phi unit distribution for all
habitats, means and 95% confidence intervals indicated. To avoid
overlap, the confidence interval bars are displaced 0.10 phi units to
the right for the Mayne Bay data and 0.05 phi units to the right for
the Trevor Channel data (Phi = -logz (sediment particle diameter)].

nant, averaging almost 60 animals/m2; Cadulus aberrans was
rarely collected.

Although the Mayne Bay and Trevor Channel sites had
similar scaphopod densities, the species assemblages were
significantly different. Pu/sellum salishorum was found in
similar abundances at all three sites. Cadulus aberrans was
absent at the Mayne Bay site and relatively abundant at the
Trevor Channel site. Dentalium rectius was about 12.5 times
as abundant as the Mayne Bay site than at the Trevor Chan-
nel site.

Buccal contents were examined from 87 Cadulus aber-
rans, 231 Dentalium rectius and 149 Pulsellum salishorum. The
proportion of each taken with food in the buccal pouch varied
substantially (Table 4). A total of 2511 items were found among
the buccal contents in C. aberrans, 654 in D. rectius and 603
in P salishorum. C. aberrans and P. salishorum buccal con-
tents consisted mainly of live, dead, or fragmental remains
of foraminiferans. D. rectius buccal contents contained a large
proportion of other items (Table 4). Both the diversity and the
evenness of the dietary array of D. rectius were higher than
in the other two species (Table 4).

Cadulus aberrans buccal contents contained 47 groups
of items, mostly foraminiferans, with foraminiferan test
fragments the third most common item (Table 4, Appendix
Table 1). Five species of foraminifera accounted for over 80%
of the total buccal contents (Appendix Table 1). The common
prey species, Cribrononion lene (Cushman and McCulloch,
1940), Elphidiella hannai (Cushman and Grant, 1927), Florilus
basispinatus (Cushman and Moyer, 1930), Rosalina
columbiensis (Cushman, 1925), and Buliminella exilis (Brady,
1884), were also well represented in the diets of Dentalium
rectius (Appendix Table 2) and Pulsellum salishorum
(Appendix Table 3). Only 1.71% of C. aberrans buccal con-
tents were other than foraminiferans.

Whole Cribrononion lene and Foraminifera fragments
dominated the buccal contents of Pulsellum salishorum, ac-
counting for almost 54% of the diet, a much larger propor-
tion as compared to Cadulus adherens (Table 4, Appendix
Table 3). Nevertheless, 29 other categories of dietary items
were also found. Non-foraminiferan food categories, e.g.
mineral grains, sediment boluses, mite eggs, and fecal pellets,
constituted 8.79% of the total buccal contents (Appendix
Table 3).

The buccal contents of Dentalium rectius included a
broader array of items. While Cribrononion lene was the most
common prey, 53 other categories of items were found (Table
4). In decreasing order, sediment particles, mite eggs, and
fecal pellets were the three next groups constituting the buc-
cal contents, cumulatively accounting for 30.89% of the diet
(Appendix Table 2). In addition to 20 species of live
foraminiferans, buccal pouch contents included substantial
diversity in other food categories: live bivalves, ostracods,
kinorhynchs, mites, barnacle cyprids, mite eggs, clear uniden-
tified eggs, turbellarians, and gastropod eggs (Appendix Table
2). Non-living dietary components included polychaete setae,
echinoid [Brisaster latifrons (A. Agassiz, 1898)] ossicles,
ostracod valves, bivalve valves, blue polypropylene rope
fragments, and several unidentified annulated objects (Appen-

SHIMEK: SCAPHOPOD DIETS 151

dix Table 2).

Infauna varied significantly among the stations, but not
seasonally within each station (Tables 5-7). Those infauna
found in the substratum were also well represented in the diets
of scaphopods, indicating that the samples were an adequate
assessment of prey availability. Several infauna taxa, par-
ticularly polychaetes, nematodes, amphipods, and harpacti-
coid copepods, were absent totally from the diet (Appendix
Tables 1-3).

The dominant prey taxon was within the protistan Order
Foraminiferida. Foraminiferans were present in the samples
from all the localities sampled and were most abundant at
the Trevor Channel site (Table 5). Foraminiferans were typically
numerically dominant at all sites, although order of abundance
varied (Tables 6, 8). Arenaceous forams, i.e. Rheophax sp.,
Saccammina sp., and Haplophragmoides sp., were typically
more common at the silty Mayne Bay and Sandford Island
sites, while overall foraminiferan species richness was greater
at the Trevor Channel station (Table 8). Living foraminiferans
were commonly eaten by all three scaphopod species (Ap-

pendix Tables 1-3) with Cribrononion lene as the most com-
mon prey of all three species of scaphopods, although its
relative proportion varied widely. Similarly, the rank order and
proportional abundances of the other five species of
foraminiferans also varied. Thus, the most common live prey
items for all three species of scaphopods studied here were
typically one of the six common foraminiferans. The only other
common living dietary items found in any scaphopod were
mite eggs that were eaten relatively frequently by Dentalium
rectius (Appendix Table 2).

An ANOVA of the arcsine transformed relative habitat
and buccal content prey taxa proportions showed that these
proportions varied significantly among predator species (Table
9). The proportions of all live prey taken by scaphopods were
compared to the proportions of those same taxa found in each
habitat and were found to differ significantly for all habitats
(Table 9). The diet and habitat proportions of the major prey
taxa were most similar at the Mayne Bay site (Fig. 3), in-
termediate at the Sandford Island site (Fig. 4), and least similar
at the Trevor Channel site (Fig. 5).

Table 3. Scaphopod abundances as determined by quantitative sediment collection.

Cadulus Dentalium Pulsellum All
aberrans rectius salishorum Scaphopods
A. Mayne Bay
Sample size 10 10 10 10
Mean + 1S. E. 0.00 + 0.00 18.00 + 4.16 6.00 + 2.67 24.00 + 4.52
B. Sandford Island
Size 8 8 8 8
Mean + 1S. E. 2.50 + 1.64 58.75 + 5.15 6.25 + 2.63 67.5 + 5.26
C. Trevor Channel
Sample size 7 7 7 7
Mean + 1S. E. 5.71 + 4.29 1.43 + 1.43 10.00 + 3.78 17.41 + 565

Table 4. Summary of all buccal pouch contents.

Cadulus aberrans

Dentalium rectius Pulsellum salishorum

Taxa Number Percent Taxa Number Percent Taxa Number Percent
A. Buccal pouch items
Live foraminiferans 28 1913 76.19 22 252 38.53 17 357 59.20
Foraminiferan tests 12 216 8.60 8 65 9.94 5 53 8.79
Other items 6 43 1.71 23 296 45.25 8 80 13.27
Foraminiferan fragments 1 339 13.5 1 4 6.27 1 113 18.74
Total 47 2511 54 654 31 603
Mean number of items 29.2 45 59
B. Proportion of scaphopods with food in buccal pouches.
C. aberrans D. rectius P. salishorum
Number examined: 87 231 149
Number with buccal contents: 86 144 102
Proportion: 0.989 0.623 0.684
C. Dietary diversity.
C. aberrans D. rectius P. salishorum
Shannon-Weiner (H’) 2.406 2.926 2.329
Evenness (J) 0.625 0.734 0.678

152 AMER. MALAC. BULL. 7(2) (1990)

a rip. fy? tT ty. i)” ey tela [at TS, ae Salle lee el Linon Gren] fern
8.8 4 8.8} a
DPS 1 | DPS |
ale 6.67 zt
6.67 4 L |
| @.4/ ‘ie aN a
P i | [
59.4 alk c |
‘dl + 10 a ‘4 aie
2 o re
0 50.2 ay =|
o o
0 fr all 5 ail
c d lanes OS C ele
ao@.2F = mins - o a a6
Ale
er +4! 7 2D oop alls |
ail: eh Le |
-0.2F |
! F
=0.32 = -O.44 a
cal AS ares MR [ge (ere ue el ere cree Ce em a Meee reer or
Bl Bx Cr El Fl Of Oa Re Bl Bx Cr El Fl Of Oa Re
FIG. 3 FIG. 4
T_T Le ae (ee | Pot | ia Ito ed Wea ing T T T
8.8 S
L ale
8.6) CDPS 4
t
|
Ce.4t |
C0.4)
I L 4
aa)
S J
0
o
q |
¢
06.2) a
| | 4
Hy
aL HH TLE |
|
-0.25 4
rare eco Tren el reel rma eer Lise ar AT

Bl Bx Cr El Fl Of 0a Rc
FIG. 5

Figs. 3-4. Proportions of prey species in the sediment and the diet; means and 95% confidence intervals indicated. The prey taxa are delimited
by the vertical dashed lines. Each prey taxon has three bars indicating, from left to right, the relative proportions of that taxon in the buccal
pouches of Dentalium rectius, D; Pulsellum salishorum, P; and the proportion of those taxa in the sediment, S. The prey taxa are: Buliminella
elegantissima, BI; B. exilis, Bx; Cribrononion lene, Cr; Elphidiella hannai, El; Florilus basispinatus, Fl; all other foraminiferans, Of; other animals,
Oa; Rosalina columbiensis, Rc. Fig. 3, Mayne Bay; Fig. 4, Sandford Island. Fig. 5. As for Figs. 3-4 except each prey taxon now has 4 vertical
bars, the farthest left bar indicating the relative proportions of that taxon in the buccal pouches of Cadulus aberrans, C. Trevor Channel.

SHIMEK: SCAPHOPOD DIETS 153

The relative mean dietary prey proportions of Dentalium
rectius and Pulsellum salishorum at Mayne Bay and Sand-
ford Island sites were not significantly different from one
another (Table 10). At the Trevor Channel site, the mean dietary
prey proportions for all three scaphopod species could be
grouped together as significantly different from the habitat pro-
portions of those same prey taxa. Alternatively, the mean prey
proportions for Cadulus aberrans, D. rectius and the habitat
form a group not significantly different from one another, but
different from the mean prey proportions found in P. salishorum
(Table 10).

Patterns of prey utilization emerged when the dietary
and habitat proportions were compared over all habitats for
each major prey species by month and habitat. Proportional
abundances of all major prey taxa or items differed significant-
ly between the habitat and the buccal contents of at least one
of the scaphopod species. These differences were consistent
and not due to seasonal or other habitat variations (Table 11).

Buliminella elegantissima was found in the gut contents
of all of scaphopods significantly less frequently than in
the associated habitats (Fig. 6). B. exilis was found in the guts
of Dentalium rectius and Pulsellum salishorum significantly less
frequently than in the associated substratum (Fig. 7). Although
the mean proportional abundance of B. exilis in Cadulus aber-
rans buccal contents was less than the habitat, the difference
was not significant (Fig. 7).

Summed over all the habitats, the mean proportional
abundances of Cribrononion lene in the substratum and buc-
cal contents of all the scaphopods did not differ significantly
(Fig. 8). In Mayne Bay site populations of Dentalium rectius
and Pulsellum salishorum; however, the mean proportional
buccal abundances of C. /ene were significantly greater than
in the habitat (Fig. 3).

Elphidiella hannai and Rosalina columbiensis were
found significantly less frequently in the guts of Dentalium rec-
tius and Pulsellum salishorum than in the sediments, while
Cadulus aberrans ate them in about the same proportion as
found in the habitats (Figs. 9, 10). A similar pattern was seen
for Florilus basispinatus; the buccal abundances were less
but not significantly so in C. aberrans (Fig. 11).

Halacaridan eggs were not eaten regularly by either
Cadulus aberrans or Pulsellum salishorum. The eggs were
taken in about the same proportion as they were found in the
environment by Dentalium rectius (Fig. 12).

There was no difference between the sizes of
foraminiferan and non-foraminiferan prey eaten by Dentalium
rectius (Fig. 13). Although D. rectius could eat items only
marginally smaller than the size of its ventral shell aperture,
the majority of the diet consisted of smaller particles and
organisms (Fig. 13).

The size-frequency distributions of the buccal contents
of Dentalium rectius (Fig. 14) and Pulsellum salishorum from
the Mayne Bay site (Fig. 15) did not differ significantly from
each other. The means of the pooled size-frequency distribu-
tions of ingested foraminiferans were significantly smaller than
those from the habitat (Fig. 16), as were the buccal contents
(Figs. 17, 18). The same is true for foraminiferans from the
Sandford Island site (Fig. 19). At both stations, relatively more

Table 5. Potential prey in Barkley Sound.

A. Abundance of foraminiferans.

Habitat Number of Foraminiferans/ml
Samples Mean + 1 Standard Error

Mayne Bay 10 0.18 + 0.08

Sandford Island 10 0.27 + 0.12

Trevor Channel 10 2.17 + 0.41

B. ANOVA - differences in the number of foraminiferans /ml of

sediment.

Source Sum of df. Mean F-ratio P
of variation Squares Square

Habitat 25.168 2 12584 19686 <0.001

small prey were ingested by scaphopods than were collected
from the habitat, nevertheless, these patterns are only subtly
different (Figs. 16, 19).

At the Trevor Channel site, all scaphopod buccal con-
tents show a preponderance of smaller prey items, the semi-
major diameter typically less than 300 um (Figs. 20-22). The
pattern of prey size utilization is similar in all three scaphopod
species. The habitat foraminiferan size frequency distribution
at this station is quite different than either of the other two
stations or the buccal contents of any of the predators (Fig.
23). Of particular interest at this station is the predominance
of predation by Cadulus aberrans in regulating the prey size
frequency distribution. C. aberrans ingested so many
foraminiferans that their cumulative distribution is effectively
that of C. abberans prey; the regulatory contributions of both
Dentalium and Pulsellum were minor.

No scaphopods studied here show a significant rela-
tionship between changes in ventral aperture width and the
size of dietary items found in the buccal pouch (Table 12). The
data are highly scattered, therefore the r-squared valules are
exceedingly low and the regressions given here represent the
best fit from several different models. The regressions were
calculated on a seasonal basis for Cadulus aberrans prey,
because of the large sample sizes (Table 12). The minor dif-
ferences in the slope and intercept of the regression lines were
not significantly different, and no line has a slope significant-
ly different from zero. None of the regressions for any
scaphopod species differed significantly from any other (Table
12).

Scaphopods processed prey at different rates. Den-
talium rectius and Pulsellum salishorum completely cleared
their buccal pouches within 36 hours. Cadulus aberrans took
over 3 days to utilize all the items in their buccal pouches
(Fig. 24). The buccal pouch contents of C. aberrans; however,
were much more numerous, and individual foraminiferans
were actually processed at a greater rate.

DISCUSSION
HABITATS

The three Barkley Sound sites differed significantly in
particle size distribution and proportion of total volatile solids,

D.

P.

Se

154 AMER. MALAC. BULL. 7(2) (1990)
—
8.391 @.45/+ a,
b Sed. Cc. a. Die Fs P. s. 4 |
Sed.
- | i
4 L
8.29} 8.351
0 | | Th) +
wi | Ww |
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z z
t | I
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FIG. 6
a al — I al
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LI es |
FIG. 8

Figs. 6-9. Mean proportional abundance of individual prey pooled for all habitats and in the buccal pouches of all the scaphopod species.
Mean abundances and 95% confidence intervals of arcsine transformed proportional abundances are shown (sediment, Sed; Cadulus aber-
rans, C.a.; Dentalium rectius, D.r.; Pulsellum salishorum, P.s.) Fig. 6, Buliminella elegantissima; Fig. 7, B. exilis; Fig. 8, Cribrononion lene; Fig.

9, Elphidiella hannai.

FIG.

SHIMEK: SCAPHOPOD DIETS 155

T T — = T
O@.4F-
Sed. Cc. a. D. r. P. s.
w 9.3-
oO |
> |
I
Qa
rs
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q 9.2 7
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FIG. 10
- ll i I
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Sed C. a. D r P. s
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s
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-@.15F- -
Po | ees ae ee
FIG. 12

although neither varied significantly from month to month
(Table 2, Fig. 1). The Sandford Island site had a significantly
higher silt fraction than did either of the other sites; the Trevor
Channel site had the smallest silt fraction. The Mayne Bay
site fraction was intermediate, but more similar to that of the
Sandford Island site.

Thomson (1981) indicated that the organic content of
the substratum along Barkley Sound ranges from about 20%

°

J

W
!

PROPORTIONAL ABUNDANCE

-8.07 r = 4

a 2 |

FIG. 11
Figs. 10-12. Mean proportional abundance of individual prey pooled
for all habitats and in the buccal pouches of all the scaphopod
species. Mean abundances and 95% confidence intervals of arcsine
transformed proportional abundances are shown (sediment, Sed;
Cadulus aberrans, C.a.; Dentalium rectius, D.r.; Pulsellum salishorum
Ps.). Fig. 10, Rosalina columbiensis; Fig. 11, Florilus basispinatus; Fig.
12, Halacaridan mite eggs.

(by weight) in the northeast to about 5% in the southwest.
The stations sampled here are consistent with that account
(Table 2, Fig. 2).

The relationship of sediment particle size distributions
and the total volatile solids found at the three stations was
complex. The proportion of coarser sediment (smaller phi
sizes) varied substantially. A consistent hierarchy was evident;
however, in sediment fractions smaller than 250 um (phi = 2).
The Mayne Bay site had more volatile solids than did the
Sandford Island site, which in turn had more than the Trevor
Channel site. Most of the particles at each station were also
smaller than 250 «m (Fig. 1), thus the trend was consistent
among stations.

The relatively high proportion of coarse organic par-
ticles at the Trevor Channel site (Fig. 2) is due to substantial
kinetic energy input during storms. As a result coarse algal
material is ground into the substratum. Observations taken
from the submersible PISCES IV confirmed large laminarian
kelp fragments on, and partially ground into, the substratum
at depths exceeding 90m. Judging by the relative paucity of
total volatile solids in the smaller particle size fractions, it is
likely these larger particles are being broken down and utilized
relatively rapidly, probably as food for infaunal organisms. The
infauna at this station were both more diverse and abundant
than at the other two sites (Tables 5, 6).

156 AMER. MALAC. BULL. 7(2) (1990)

Table 6. Infauna collected.

A. MAYNE BAY
Month/Year 12/83 12/83 3/84 3/84 6/84 6/84 9/84 9/84 12/84 12/84 Total
Number of samples 1 2 1 2 1 2 1 2 1 2 collected
Volume (ml) 35 40 40 61 93 71 59 56 58 56.5
FORAMINIFERA
Astrorhizidae sp. 1 1
Buliminella elegantissima 3 3
B. exilis 2 7 4 13
Cribrononion lene 8 6 1 2 1 3 5 26
Elphidiella hannati 1 1 1 3
Florilus basispinatus 17 5 1 23
Lagena sp. A 1 1
Nonionella stella 1
Rheophax sp. 1 1
Rosalina columbiensis 2 4 2 8
Saccammina 1 2 3
OTHER INVERTEBRATES
Mite eggs 3 1 2 1 7
Kinorhynch sp. 2 2
Polychaete sp. 6 6
Amphipod sp. 1 1
Nematode sp. 1 1
Harpacticoid sp. 2 4 6
Axinopsida serricata 10
Ophiuroid sp. 1 1
Ostracod sp. 1 1
TOTAL 37 23 1 4 5 4 0 2 27 15 118
B. SANDFORD ISLAND
Month/Year 12/83 12/83 3/84 3/84 6/84 6/84 9/84 9/84 12/84 12/84 Total
Number of samples 1 2 1 2 1 2 1 2 1 2 Collected
Volume (ml) 57 65 58 44 69 85 54 62.5 59 58.5
FORAMINIFERA
Astrorhizidae sp. 1 3 4
Buliminella elegantissima 15 24 39
Cibicides sp. 1 1
Cribrononion lene 2 3 2 3 10
Elphidiella hannai 1 1
Florilus basispinatus 2 1 5 4 7 19
Globobulimina 2 1 9 6 17
Haplophragmoides sp. 1 1
Hippocrenella sp. 1
Lagena sp. A 1 2
L. sp. B 1 1
L. sp. D 1 1
L. sp. E 2 2
Nonion sp. 3 4 11 18
Nonionella stella 2 2 7 11
Rosalina columbiensis 4 1 4 9
Rheophax sp. 6 1 7
Rotorbinella sp. 2 2
Saccammina sp. 2 5 4 11
Spirulina sp. 1 2 2 iS)
Textularia sp. 2 2
Triloculina sp. A 1 1
OTHER INVERTEBRATES
Mite eggs 1 1 1 2 2 1 8
Kinorhynch sp. 1 4 5
Ostracod sp. 1 2 2 5
Polychaete sp. 1 1
Nematode sp. 2 1 3 100 106
Corophium sp. 1 1
Harpacticoid sp. 6 6
297

TOTAL 3 2 7 7 11 10 8 8 62 179

SHIMEK: SCAPHOPOD DIETS 17

Table 6. (continued)

C. TREVOR CHANNEL

Month/Year 12/83 12/83 3/84 3/84
Number of samples 1 2 1 2
Volume (ml) 59 65 56 52

6/84 6/84 9/84 9/84 12/84 12/84 Total

1 2 1 2 1 2 Collected
59 69 57 49.5 52.5 39.8

FORAMINIFERANS
Astrorhizidae sp. 2
Astrononion sp.
Buliminella sp. C
B. sp. D
B. elegantissima
B. exilis
Cibicides sp.
Cribrononion lene 3 2
Discorbinella sp.
Elphidiella hannai 6 8 37 47
Faujacina sp. 5
Florilus basispinatus 144 72 34 26
Globobulimina sp. 8 4 4 2
Haplophragmoides sp. 3
Lagena sp. A 1
L. sp. B 1
L. sp. C 1
L. sp. D
Nonion sp. D
Nonionella stella
Quinqueloculina sp.
Rosalina columbiensis 1 2 5
Rheophax sp.
Rotorbinella sp.
Saccammina sp.
Spirulina sp. 1
Textularia sp.
Triloculina sp. A 1 4 3
T sp. B 2 1
T sp. C
T sp. D
Unidentified Foraminiferan
Uvigerina sp. 1

OTHER INVERTEBRATES
Axinopsida serricata
Compsomyax subdiaphana 2 2
Mytilus sp.
Mite eggs 3
Kinorhynch sp.
Ostracod sp. 2 2
Harpacticoid sp.
Nematode sp.
Tanaid (Leptochelia?)
Corophium sp.

TOTAL 162 93 93 95

=

=

70 2 20 99 12 301

60 60 15 15 57 30 513

wo
a
=
foe)
-a+-+++NhM-Of 0 Wwwn-fbn
w ine) epee WwW
= = (oy) NO
O- + ONDODMD-]ANWO-OD-LHW

ine)
—-~ A
£
N
Oo

71 167 71 57 268 216 1293

Scaphopod abundances sampled here were much
higher than hitherto reported from other areas (Gainey, 1972;
Bilyard, 1974). These high abundances are not rare; however,
dredging reports from other fjord systems on Vancouver Island
indicate similar scaphopod abundances in deeper areas (W.
Austin, pers. comm.). High scaphopod densities probably oc-
cur in all silty fjord environments of Northern British Columbia
and Southeastern Alaska.

It is likely that the scaphopod abundances documented
here (Table 3) reflect substantial underestimates of total abun-
dances. These data were based on samples taken by Petersen
grabs, which typically penetrate the bottom only to a depth
of 15 to 20 cm (Holme and Mcintyre, 1971). Laboratory obser-

vatons made here and observations by Poon (1987) and J.
Levitt (pers. comm.), confirm that the scaphopods studied
here are capable of burrowing to a depth of up to 30 to 40
cm in aerobic substrata (Shimek, 1988, 1989). The depth of
the redox discontinuity is unknown from stations studied here,
but exceeds the sampling depth; no samples had indication
of anaerobiosis. | believe actual scaphopod densities to be
two to five times higher than these reported here.

INFAUNA

The sample size for the infaunal examination was ade-
quate to determine abundances of common taxa at the Mayne
Bay and Sandford Island sites, but was less reliable in regard

158 AMER. MALAC. BULL. 7(2) (1990)

to uncommon taxa. Sample sizes at the Trevor Channel site
were sufficiently large to determine all infaunal abundances.

No seasonal trends in infaunal abundances could be
demonstrated at any station (Tables 6, 7), although this could
be an artifact of the variance introduced by relatively small
foraminiferan sample sizes from the Mayne Bay and Sand-
ford Island sites. However, no statistically significant changes
were found at the Trevor Channel site.

Infaunal abundances did differ significantly among the
three sites. This is best seen in the foraminiferan abundances
although similar trends in other taxa can be seen as well
(Tables 5, 6, 7, 8). Although the foraminiferans were general-
ly dominated by the same group of species at all stations, the
rank order and proportional abundances did vary.

Differences in prey abundances can be related to
scaphopods in one of two ways: 1) scaphopods passively
tracked prey populations with regard to their diets, which
would be evident if they had the same relative proportions
of any given prey in their guts as were found in the native
substratum; 2) scaphopods could be altering the distributions
of their prey. The latter condition would be supported if there
was evidence of active selection or rejection of individual prey.

POTENTIAL PREY

The ANOVA on the proportional abundances of the
common prey were sufficiently robust to demonstrate signifi-
cant variation in the foraminiferan abundance among habitats
(Table 5). In addition, the major prey taxa abundances differed
significantly in all of the habitats, but not from month to month,
or between the taxa seasonally (Table 7). The pattern of varia-
tion and the significance of the interactive terms was similar
from all three areas, indicating the samples tracked consis-
tent patterns throughout the Barkley Sound area.

Previous studies (Lacaze-Duthiers, 1856, 1857; Mor-
ton, 1959; Pilsbry and Sharp, 1897-98, Dinamani, 1964; Fisher-
Piette and Franc, 1968; Gainey, 1972; Bilyard, 1974; Carter,
1983; Poon, 1987) indicated the major prey of scaphopods
were foraminiferans. Consequently, | examined the variation
in foraminiferan abundances in detail. At all sites
foraminiferans numerically dominated the infaunal com-
munities. Although organisms smaller than 63 »m were
regularly found in scaphopod gut contents, substrata
examined for infauna were sieved with a 63 um screen to
remove silt and clay. Therefore, potential prey size-frequency
data are reliable only for size classes greater than 63 um.
Organisms less than 10 »m in diameter were not analyzed
and it is likely bacteria and small eukaryotic organisms were
quite abundant. These organisms comprise food for
foraminiferans, and contribute directly to the diet of the
deposit-feeding Dentalium rectius. Abundances of micro-
organisms can be only inferred.

DIETS

Previous observations on scaphopod diets have been
based either on small data sets generated from relatively few
individuals from one population (Gainey, 1972; Bilyard, 1974;
Taib, 1980; Poon, 1987) or various species (Carter, 1983). With

the exception of Bilyard (1974) and Poon (1987), the taxonomic
precision of the dietary determinations has been inadequate
for detailed analysis. Bilyard (op. cit.) recognized selection
and rejection of potential dietary items by Dentalium entale
stimpsoni Henderson, 1920. Poon (op. cit.) also found
restricted diets in Cadulus tolmiei. The diets reported here are
consistent with their observations: the scaphopods studied
herein selectively accept or reject individual food items
(Shimek, 1988).

The prey collected from scaphopod buccal pouches
represented a diverse array of whole organisms and other
items. All scaphopods preyed on foraminiferans. However, the
relationships of these and other prey taxa varied significant-
ly among the predator species (Table 4, Appendix Tables 1-3).
Additionally, while buccal contents differed significantly from
area to area, seasonal variations were insignificant (Figs. 3-5;
Table 9).

CADULUS ABERRANS

This species fed in accordance with stereotypical
scaphopods, specializing on live foraminiferans which were
numerically dominant, although empty foraminiferan tests and
test fragments also comprised a substantial fraction of the
prey (Table 4). Cadulus aberrans also fed more frequently:
98.9% of individuals had prey in the buccal pouches, and had
more food items in their buccal pouches (mean = 29.2) than
did either of the other species. Total prey number could be
high: one individual had 135 items in its buccal pouch.

The size of the predator, as measured by ventral aper-
ture width, was not related to the size of the prey. No signifi-
cant changes in the sizes of the buccal contents occurred with
the seasons.

Prey composition varied; however, five species of live
foraminiferans and foraminiferan fragments comprised over
80% of the diet (Appendix Table 1). Dietary diversity was low
(H’ = 2.406) but higher than that of the other foraminiferan
predator Pulsellum. The evenness index (J = 0.625) was the
lowest of all scaphopods studied indicating the numerical
dominance of those few prey taxa (Table 4).

PULSELLUM SALISHORUM

This species also preyed mostly upon live
foraminiferans, although dead foraminiferan remains con-
stituted a substantially larger component of their diets than
in Cadulus aberrans. Live Cribrononion lene dominated the
diet with empty foraminiferan tests next most common.

Foraminiferan tests, like other particulate mineral mat-
ter in benthic ecosystems, become colonized by bacteria.
These tests, therefore, can be desirable food sources; bacteria
have a relatively high nitrogen to carbon ratio (Dales, 1964;
Meadows, 1964). In addition, the tests could be an important
dietary source at calcium carbonate for scaphopods in silty
environments.

About 42% of the total buccal contents, 256 items, con-
sisted of live Cribrononion lene. The rest of the seven most
common dietary categories were dead items and together with
C. lene represented over 80% of total buccal contents (Ap-
pendix Table 3). Selection for a single prey type was reflected

SHIMEK: SCAPHOPOD DIETS 159

Table 7. Analysis of variance to test for differences in proportional prey abundances.

Source of variation Sum of df. Mean F-ratio P

squares square

MAYNE BAY
MAIN EFFECTS 4.526 10 0.453 5.506 < 0.0001
Prey taxon 4.342 i 0.620 7545 < 0.0001
Month sampled 0.197 3 0.066 0.797 0.5001
2-FACTOR INTERACTIONS
Taxa by months 1.861 21 0.089 1.078 0.3940
RESIDUAL 5.014 61 0.082
TOTAL 11.401 92
SANDFORD ISLAND
MAIN EFFECTS 5.645 10 0.564 4.282 0.0001
Prey taxon 5.530 7 0.790 5.992 < 0.0001
Month sampled 0.095 3 0.032 0.241 0.8678
2-FACTOR INTERACTIONS
Taxa by month 2.093 21 0.100 0.756 0.7600
RESIDUAL 9.229 70 0.132
TOTAL 16.968 101
TREVOR CHANNEL

MAIN EFFECTS 4.891 10 0.489 9.632 < 0.0001
Prey taxon 4.859 ‘Gi 0.694 13.670 < 0.0001
Month sampled 0.036 3 0.012 0.239 0.8690
2-FACTOR INTERACTIONS
Taxa by months 1.038 21 0.049 0.973 0.4995
RESIDUAL 7.464 147 0.051
TOTAL 13.390 178

in the Shannon-Wiener Index (H’ = 2.329), which was lower
for this species than the other two scaphopods studied here.

Only 68.4% of Pulsellum salishorum had buccal pouch
contents which averaged 5.9 prey items per individual, far
fewer than in the other foraminiferan specialist, Cadulus aber-
rans, and more than Dentalium rectius (Table 4). No seasonal
or habitat patterns in feeding were seen. Nor were any pat-
terns evident regarding the relative sizes of predator and prey.
P salishorum was about equally abundant in all three habitats
sampled. This could reflect the lack of foraminiferan prey at
the Mayne Bay and Sandford Island sites, and competition
from the more active and voracious C. aberrans at the Trevor
Channel site.

DENTALIUM RECTIUS

Only 62% of Dentalium rectius contained food in the
gut, averaging 4.54 prey items per individual. The diet was
also more evenly distributed among the other categories of
prey items (J = 0.734), as compared to Cadulus aberrans and
Pulsellum salishorum (Table 4). Again, no significant patterns
relating predator and prey sizes were evident, probably due
to the high variability in prey.

Although Cribononion lene was the most common live
prey organism, most of the buccal pouch contents did not con-
sist of live foraminiferans (Table 4, Appendix Table 2, Fig. 13).
Sediment, compacted into small boluses, was also commonly
ingested, as were fecal pellets, mineral sediment grains, and
foraminiferan fragments. The surfaces of these items could

be sources of bacteria which probably constitute a major food
source for this species. Sediment and mineral grains have
been noted in the diets of Dentalium entalis L. (1980) and D.
stimpsoni (Bilyard, 1974), although little significance has been
attached to these items as sources of nutrition. Bright gold-
colored eggs presumed, by comparison, to be halacarid mite
eggs, were the second most commonly ingested food item
of Dentalium rectius.

Dietary diversity was greater (H’ = 2.93) than that of
either of the other two species of scaphopods. Of the
organisms collected from the substrata, only polychaetes,
nematodes, and harpacticoid copepods were not found in the
gut of at least one specimen of Dentalium rectius. It is likely
these animals move rapidly or vigorously enough to avoid cap-
ture by captacular attachment.

The captacular morphology of Dentalium rectius allows
collection of fine particulate material. This mode of feeding
is well documented among dentalioid scaphopods (Dinamani,
1964; Gainey, 1972; Bilyard, 1974; Shimek, 1988). The wide
array of dietary items eaten by D. rectius reflects the effec-
tiveness of this feeding mode. Several types of very fine par-
ticulate matter were commonly in the form of boluses in the
buccal pouches. In addition, many of the mineral grains,
foraminiferan fragments, small uniloculate foraminiferans, and
unidentified black spherules (diameters to 30 4m) could also
have been collected by the captacular ciliary band.

Many of the prey of Dentalium rectius were not capable
of rapid or sustained motion, as were virtually none of the prey

160 AMER. MALAC. BULL. 7(2) (1990)

T T T T T T a T T T T | i T i T 7 T |
9.285 a
NON-FORAMINIFERANS
N = 387
@.18F 7
>
oO
Z
5
a: 68 a
Wu
o&
Te
i)
=> mom
H
ke a
a
al
WW
OY
@.12+ FORAMINIFERANS
N = 255
@.22F 7
| do 4 L | A 4 1 | 1 1 1 | nail 4 _L | elt L |
8 200 400 600 800 1000
Size (um)

Fig. 13. Size frequency distribution of Dentalium rectius buccal con-
tents, pooled over all habitats. The mean + 1 standard deviation for
the non-foraminiferan distribution = 170 + 116um, for the foramini-
feran distribution = 174 + 103 um. The computed t statistic for the dif-
ference in the means: t = -0.419; P = 0.675, not significant; « = 0.05.

| T 7 T TT T | T T a ] qi T T TT T T
6.57 4
Oieret s= N = 96 a

L 4

+ 8.37 =
[a] | |
aes
W 4
=]
Go f 4
Ww |
Y }
w 8.2) +
re 4
VA |
|
LU 4
Q.4 Ly |
| a |
a | VG = =
oe ees rie | a | 1 1 | L 1 L | 1 1 1 |
(2) 2068 400 680 800 10080

Size (pum)

Fig. 15. Mayne Bay. Size frequency distribution of Pu/lsellum
salishorum buccal contents.

T i T T T T T T ¥ mi T T T T T T T T
6.2) 4
6.16>- N = 382 4
>+8.12>- =|
Oo
> |
Ww
=) bE
Go
W 4
aX
Ug.as- +
3 al
|
6.04 >- =|
+ 4
Ca 4
| a =! 1 | 1 2 1 | 4 1 1 | 4 4 1 | L a L |
(2) 200 400 600 8e0 1600

SIZE (pm)

Fig. 14. Mayne Bay. Size frequency distribution of Dentalium rectius
buccal contents.

if T T T 1 T T T T T T | T T T
@.27> 4
BUCCAL CONTENTS
N = 197
@.17> .
> |
oO
Zz.
5
39-87 4
w
©
uw
w
>
H |
+ 8.863>- a
I
a
i
a
@.13> S
SEDIMENT
N = 166
0.23 4
L a ee Pree Lae ae ear el (ese
() 208 400 600 800
SIZE (um)

Fig. 16. Mayne Bay. Size frequency distribution of sediment and buc-
cal content foraminiferans. Buccal content foraminiferans are pooled
over all Scaphopods. The mean + 1 standard deviation for the sedi-
ment foraminiferan distribution = 245 + 143 um; for the buccal
foraminiferan distribution = 142 + 93 um. The computed t statistic
for the difference in means: t = -8.226; P = 9.922 x 10-9, highly signifi-
cant; a = 0.05.

SHIMEK: SCAPHOPOD DIETS

96

\\
SS

KAXQQQ QQ ggg ggg gaa QQ GJ KF

ee

KS

8.25

97

KKK,,-D MK5[J5K GK ——

D0YW

CN
=

—
.W

=
V.D.VV YY
KK = Se

= ——

es

8.15);

ASNANDAYS

8. 06-
8.035

400 680

200

400 680 8oe 1608

200

Size (pum)
Fig. 17. Sandford Island. Size frequency distribution of Dentalium rec-

tius buccal contents.

Size (pum)

Fig. 18. Sandford Island. Size frequency distribution of Pulsellum

salishorum buccal contents.

0.335

8.23F

Oe. 13+

SNanDaas anil laa

8.17)

8.275

f-
o
ral
a
iT)
z
=
=
es KK KR—hj#(( cen
AWC iQ GG G KE. »» = A\VJJV0V.\VV
ee SS
.)V.YVCWO ee A aEEEEnEeaaenEe AY
WWW KWKCKWKKKG GCC GK oes cee enn monn
MK KK —>p_oQycyyvv“v SS
== | =a l 1 ae 1 |
©
ee oe
a = fan) fo) fa) i
a ° AONaNDAaYS as
T fav)
fav)
wo
rm)
a onl
Z Ul
a
3 ' 8
a) 5 Fb 2
t Z ©
41 0 “9
Io =: Hod ~
oO SN o " €
ou “q 5
5 Ww
m Zz ‘ i Oo Z oon
OS tw
N
H
o
fax)
[ux]
cu
fax)

Fig. 19. Sandford Island. Size frequency distribution of sediment and
buccal content foraminiferans. Buccal content foraminiferans are

pooled over all scaphopods. The mean + 1 standard deviation for

the sediment foraminiferan distribution

400 600 860 1990

2060

Cum)

Size
Fig. 20. Trevor Channel Site. Size frequency distribution of Cadulus

aberrans buccal contents.

0.0377,

; for the buc-

-2.090; P

190 + 128 um

0.05.

cal foraminiferan distribution = 154 + 134 um. The computed t

statistic for the difference in the means: t

significantly different; a

162 AMER. MALAC. BULL. 7(2) (1990)

@.18F =]
Zé |
-
Hp
6.15 AD N = 162
gee
a
|
La
@.12 Ll =
> L
Z VA yy
46.09 WH |
é HH
Y Ha
L lille
0.06 a
a
| a
my
0.03>- | cee
| ii
| : i
9 | Ge HEL ol
) 260 400 680 8ea 1008

Size (um)

Fig. 21. Trevor Channel Site. Size frequency distribution of Dentalium
rectius buccal contents.

Lees
2
S
N = 293
VA |
6.3) :
i
; :
(a) y
co | | a |
oo
iL Li
VA
Yio |
8.1} 17
[Eg
a. i=
Q | wm 7 G} Vz A dacccmmzz, ZA ZN —
i L 1 " a tt it n 1 i J . 1 ia it He)
) 200 400 60e 800 1000
Size (um)

Fig. 22. Trevor Channel Site. Size frequency distribution of Pulsellum
salishorum buccal contents.

BUCCAL CONTENTS

N = 2027

SEDIMENT
N = 1260
9.23) J
eal ae ns Sr | 1 , | 1 L | n = |
(2) 300 680 900 1200 1500
SIZE (um)

Fig. 23. Trevor Channel Site. Size frequency distribution of sediment
and buccal content foraminiferans. Buccal content foraminiferans are
pooled over all scaphopods. The mean + 1 mean standard devia-

tion for the sediment foraminiferan distribution = 368 + 177 um; for
the buccal foraminiferan distribution = 163 + 106 um. The computed
t statistic for the difference in the means: t = -41.419; P = 1.010 x
10-7, highly significant; a = 0.05.

of either Cadulus aberrans or Pulsellum salishorum. Never-
theless, at least occasionally, a few relatively mobile prey were
caught. These included ostracods, a barnacle cyprid, a mite,
and several kinorhynchs. Their susceptibility to predation
could be caused by properties of their cuticles or their lack
of vigorous directed locomotion.

PREY SPECIALIZATION BY TAXON

All three species ingested items that appear to have
little nutritive value. Bilyard (1974) found that Dentalium entale
stimpsoni ate few empty foraminiferan tests. By calculating
electivities he (Bilyard, op. cit). concluded empty tests were
not desired food items. Empty foraminiferan tests and test
fragments are found commonly in the sediment and in the
buccal contents of all three species of scaphopods examined
here.

It is possible to assess selectivity of predation within
the foraminiferan component of the total dietary array by com-
paring proportional prey abundances. If the proportional abun-
dances found in the buccal pouches approximated the pro-
portional abundances for the same taxon in the native habitat,
then the scaphopods were harvesting the foraminiferans as
they were encountered. If the abundances in the buccal

SHIMEK: SCAPHOPOD DIETS 163

pouches were greater than in the habitat, the scaphopods
were presumably actively selecting prey items. If the abun-
dances of the foraminiferans in the gut were less than found
in the native substratum, the scaphopods were presumably
actively rejecting these potential prey.

The most abundant foraminiferans found in the
scaphopod buccal pouches were also among the most com-
mon foraminiferan taxa in the habitat, although in some cases
other species were more abundant (Appendix Tables 1-3,
Tables 6-8). All foraminiferans, and the halacarid mite eggs,
were designated as ‘‘potential food items’’ and their abun-
dances were examined at each habitat on a seasonal basis.
These abundances were significantly different from one
another, but the relative proportional abundances did not vary
significantly among seasons in any habitat examined (Table
7). Because these taxa did not vary significantly seasonally,
the data were pooled over all the sampling periods, and the
proportional abundances of these taxa were calculated (Table
8). Pooling had the effect of increasing the effective sample
size for the Mayne Bay and Sandford Island sites infauna,
possibly ameliorating the problems of foraminiferan rarity.

The ANOVA of the relative prey proportions in the
habitat or buccal contents had three components (Table 9).
The first test indicated significant differences among the
potential prey taxa; they were not equally abundant. The se-
cond test indicated, except at the Trevor Channel site, signifi-
cant differences in the pooled abundances of potential prey
from the habitat and buccal contents. The two-factor interac-
tion tested for significant differences in the relative abun-
dances of foraminiferans between the predators’ diets and
the sediment for each habitat. In all cases the two-factor tests
showed highly significantly differences in proportional abun-
dances (Table 9).

The pattern of these proportional abundance dif-
ferences was similar for all habitats (Table 10). At Mayne Bay
and Sandford Island sites, Dentalium rectius and Pusellum
salishorum had mean proportional prey abundances that were
not significantly different from one another, but the relative
proportion of potential prey was significantly less in their buc-
cal contents than in the habitats; the predators were not in-
gesting foraminiferans at a frequency equal to the number
of encounters. At the Trevor Channel site, all three scaphopods
followed a similar pattern; however, the mean relative propor-
tional abundances of each foraminiferan species found in the
substratum was lower. This was likely an artifact of the
increased foraminiferan diversity at this site, coupled with the
lack of dominance of any one species. Thus, typically any
potential foraminiferan prey was part of a larger species ar-
ray than at the other two stations, and constituted a propor-
tionally smaller fractional component of the fauna. The relative
proportional abundances of Cadulus aberrans and D. rectius
prey items, and the potential prey from the habitat are not
significantly different from one another. Habitat and potential
prey abundances do differ for P salishorum. Likewise prey
abundances did not differ significantly among the three
scaphopods but they did differ from the respective sediment
abundances (Table 10).

Therefore, at the Trevor Channel site, the lower mean

Table 8. Habitat foraminiferan abundances as a proportion of the total
enumerated prey.

Species Mayne Sandford Trevor
Bay Island Channel

Astrononion sp. — — 0.011
Astrorhiza sp. 0.004 0.019 0.011
Buliminella elegantissima 0.049 0.154 0.018
B. exilis 0.228 0.142 0.102
B. sp. C — — 0.011
B. sp. D — — 0.011
Cibicides sp. — 0.013 0.013
Cribrononion lene 0.227 0.067 0.053
Discorbinella sp. — — 0.010
Elphidiella hannai 0.017 0.012 0.134
Faujacina sp. — _ 0.017
Florilus basispinatus 0.196 0.079 0.223
Globobulimina sp. 0.012 0.069 0.038
Haplophragmoides sp. — 0.003 0.013
Hippocrenella sp. — 0.004 —

Lagena sp. A 0.007 0.008 0.011
L. sp. B — 0.003 0.011
L. sp. C — — 0.011
L. sp. D — 0.013 0.011
L. sp. E _— 0.007 _

Nonion sp. D — 0.066 0.012
Nonionella stella 0.016 0.127 0.024
Quinculoculina sp. — — 0.011
Rheophax sp. 0.051 0.022 0.014
Rosalina columbiensis 0.124 0.076 0.025
Rotorbinella sp. — 0.007 0.013
Saccammina sp. 0.009 0.039 0.015
Spirillina sp. — 0.030 0.042
Textularia sp. 0.012 0.037 0.028
T. sp. B — -- 0.012
Triloculina sp. A 0.012 0.004 0.021
T sp. B — — 0.014
T sp. C — = 0.013
T sp. D — - 0.011
Uvigerina sp. _— — 0.021
Unidentified 0.037 _ 0.014

proportional abundances of potential prey in the sediment
coupled with the diverse diet of Cadulus aberrans make the
differences less distinct. However, the mean proportional
abundances of foraminiferans in buccal contents of the three
scaphopods are significantly lower than those found in the
sediment, a pattern identical to those of the other two areas
(Table 10).

This prey utilization pattern indicates that scaphopods
typically rejected most of the potential food items that they
encountered. Detailed examination of the six most abundant
foraminiferans found in the buccal pouches, as well as utiliza-
tion of halacarid mite eggs, reveals different patterns for the
utilization of each major prey taxon (Table 11).

Buliminella elegantissima was eaten less frequently
than expected by all three scaphopod species (Table 11, Fig.
6). A different pattern was shown by B. exilis and Florilus
basispinatus, which were preyed upon significantly less fre-
quently by Dentalium rectius and Pulsellum salishorum.

164 AMER. MALAC. BULL. 7(2) (1990)

Table 9. Comparison of the arcsine transformed prey proportional abundances from the habitat and scaphopod

buccal contents.

Source of variation Sum of df. Mean F-ratio P
squares square
MAYNE BAY

MAIN EFFECTS 5.678 9 0.631 10.407 <0.0001
Prey taxon 3.461 7 0.494 8.156 <0.0001
Source (habitat or buccal

contents by species) 1.349 2 0.674 11.123 0.0001
2-FACTOR INTERACTIONS
Taxa by source 1.541 14 0.110 1.815 0.0535
RESIDUAL 4.183 69 0.061
TOTAL 11.402 92

SANDFORD ISLAND

MAIN EFFECTS 6.515 9 0.724 8.576 <0.0001
Prey taxon 3.819 7 0.546 6.463 < 0.0001
Source (habitat or buccal

contents by species) 1.093 2 0.546 6.472 0.0026
2-FACTOR INTERACTIONS
Taxa by source 3.039 13 0.234 2.769 0.0031
RESIDUAL 5.993 71 0.084
TOTAL 15.547 93

TREVOR CHANNEL

MAIN EFFECTS 5.002 10 0.500 16.945 < 0.0001
Prey taxon 4.636 7 0.662 22.437 <0.0001
Source (habitat or buccal

contents by species) 0.147 3 0.049 1.661 0.1779
2-FACTOR INTERACTIONS
Prey and source 4.052 21 0.193 6.538 < 0.0001
RESIDUAL 4.339 147 0.030
TOTAL 13.393 178

Although the mean proportions of these prey species in the
buccal contents Cadulus aberrans were less than expected,

Table 10. Multiple range tests of the pooled prey abundances as a
proportion of the total potential prey abundances from the sediment
and the buccal contents of each scaphopod species examined.
Homogeneous groups, indicated by the same letter, do not have
significantly different mean proportional prey abundances.

Source Mean
proportional
prey abundances

Homogeneous
groups

MAYNE BAY

Pulsellum salishorum 0.224 A
Dentalium rectius 0.259 A
Sediment 0.573 B

SANDFORD ISLAND

P. salishorum 0.218 A
D. rectius 0.255 A
Sediment 0.599 B

TREVOR CHANNEL

P. salishorum 0.255 A
D. rectius 0.283 A B
Cadulus aberrans 0.370 B

the difference was not statistically significant (Table 11, Fig.
7, 11).

Elphidiella hannai and Rosalina columbiensis were in-
gested by Cadulus aberrans about as frequently as they were
encountered. Both of these foraminiferans were ingested
significantly less frequently by Dentalium rectius and Pulsellum
salishorum (Table 11, Figs. 9, 11).

The mean proportion of Cribrononoin lene in buccal
contents was higher than that in the sediment for all three
scaphopod species, although the elevation for Dentalium rec-
tius was minimal. None of the elevations was significant if the
data were pooled over all the habitats (Table 11, Fig. 8). Oc-
casionally in some habitats, the elevation was significant
(Fig. 5).

Halacaridan egg predation provided an interesting con-
trast to that of foraminiferans (Fig. 12). The eggs were ingested
slightly less frequently than they were encountered by Den-
talium rectius. The eggs were rarely eaten by the other two
scaphopods species. This difference was highly significant
(Table 11).

Sufficient data were available to test for prey utilization
differences on a seasonal basis for all major taxa except for
Buliminella sp. No significant differences were found.

Some differences in proportional usage occurred be-
tween the habitats. For example, while pooled data indicated

SHIMEK: SCAPHOPOD DIETS 165

Table 11. Analysis of variance for major prey arcsine transformed proportional abundances.

Source of variation Sum of df. Mean F-ratio P
squares square
Buliminella elegantissima
MAIN EFFECTS
Category (habitat, diet) 0.346 3 0.115 6.892 0.0009
RESIDUAL 0.603 36 0.017
TOTAL 0.949 39

Buliminella exilis
MAIN EFFECTS

Category (habitat, prey) 0.379 3 0.126 4811 0.0066
RESIDUAL 0.919 35 0.026
TOTAL 1.298 38

Cribrononion lene
MAIN EFFECTS 0.398 6 0.066 1.246 0.3088
Category (habitat, prey) 0.237 3 0.079 1.480 0.2379
Month 0.182 3 0.061 1.136 0.3489
2-FACTOR INTERACTIONS
Category by months 0.211 9 0.023 0.440 0.9033
RESIDUAL 1.759 33 0.053
TOTAL 2.368 48

Elphidiella hannai
MAIN EFFECTS 1.095 6 0.182 6.477 0.0002
Category (habitat, diet) 1.030 3 0.343 12.197 0.0000
Months 0.129 3 0.043 1.524 0.2293
2-FACTOR INTERACTIONS
Category by months 0.275 9 0.031 1.085 0.4029
RESIDUAL 0.817 29 0.028
TOTAL 2.186 44

Florilus basispinatus
MAIN EFFECTS 2.973 6 0.495 7.011 0.0001
Category (habitat, prey) 2.363 3 0.787 11.142 < 0.0001
Months 0.107 3 0.036 0.506 0.6811
2-FACTOR INTERACTIONS
Category by monthws 0.199 9 0.022 0.313 0.9652
RESIDUAL 2.332 33 0.071
TOTAL 5.504 48

Rosalina columbiensis
MAIN EFFECTS 0.358 6 0.060 3.183 0.0150
Category (habitat, prey) 0.295 3 0.098 5.237 0.0048
Months 0.40 3 0.013 0.703 0.5572
2-FACTOR INTERACTIONS
Category by months 0.285 9 0.032 1.689 0.1341
RESIDUAL 0.582 31 0.019
TOTAL 1.225 46

Halacarid eggs
MAIN EFFECTS 1.201 6 0.200 6.969 0.0001
Category (habitat, prey) 1.097 3 0.366 12.736 <0.0001
Months 0.095 3 0.032 1.102 0.3647
2-FACTOR INTERACTIONS
Category by prey 0.326 9 0.036 1.261 0.3005
RESIDUAL 0.804 28 0.029

TOTAL 2.331 43

166 AMER. MALAC. BULL. 7(2) (1990)

that Cribrononion lene were eaten more frequently than ex-
pected, utilization rates were not significantly different between
the habitat and the buccal contents. At the Trevor Channel
site, however, all three species of scaphopods ate C. /ene
significantly more frequently than expected (Fig. 5).

PREY SPECIALIZATION BY SIZE

All three scaphopods ingested prey up to about
1.00 mm in semimajor axis diameter. The upper limit of prey
size was effectively the ventral shell aperture diameter. The
preponderance of all items ingested however, was less than
300 »m in diameter.

The ingested prey size-frequency distributions were
consistent within each species across the habitats. The ma-
jority of the prey eaten by Dentalium rectius were smaller than
200 um, nevertheless, a substantial portion of its diet was com-
posed of larger items (Fig. 13, 14, 17, 21). The sizes of the
foraminiferan and non-foraminiferan prey items did not differ
significantly (Fig. 13).

Pulsellum salishorum ate smaller prey than the other
scaphopods; most of its prey were smaller than 100 um in
diameter (Figs. 15, 18, 22), although prey size varied. P
salishorum is smaller than Dentalium rectius (Shimek, 1989),
and the slight difference in the respective buccal content size-
frequency distributions could be a reflection of this disparity.
There were no significant increases in the size of the buccal
contents with increases in the size of the ventral aperture of
the scaphopod shell (Table 12).

The median prey size was taken by Cadulus aberrans
at Trevor Channel site was between that of the other two
species, although the total ranges broadly overlapped. Par-
ticularly between the two selective foraminiferan predators;
however, the minor differences in the median sizes of prey
eaten could be important in facilitating differential prey utiliza-
tion. The mean adult ventral aperture width between these
species did not differ (Shimek, 1989).

EFFECTS OF PREDATION

Without the results of experimental manipulation
(Shimek, unpub. data), unambiguous determination of the
result of scaphopod predation on the infauna is impossible.
Nonetheless, circumstantial evidence suggested that signifi-

Table 12. Regression analysis of ventral aperture width vs. prey size,
linear model: Y = a+bX, no regression has a slope significantly
different from 0.

R-squared Number

Cadulus aberrans

March: Y = 122.13 + 56.313X 0.35 1282

June: Y = 195.30 — 48.347X 0.42 234

Sept.: Y = 291.69 — 128.198X 5.45 143

Dec.: Y = 10383 + 9.876X 0.02 714
Dentalium rectius

All: Y = 104.25 + 33.925xX 3.04 642
Pulsellum salishorum

All: Y = 14.672 + 83.296X 1.97 485

cant effects are caused by this guild of infaunal predators.
Both the mean sizes, and the size-frequency distributions of
cumulative foraminiferan prey eaten, were significantly dif-
ferent from the mean sizes and size-frequency distributions
of the foraminiferans collected from the habitat. This was
especially notable at the Trevor Channel station where
Cadulus aberrans seemed to exert substantial predation
pressure on infaunal foraminiferans.

Although all three scaphopod species were found at
this station, the number of the foraminiferans eaten by Cadulus
aberrans was substantially greater. Also, processing of in-
gested prey occurred more rapidly in C. aberrans (Fig. 24).
Nevertheless, the predatory effect was cumulative among all
three scaphopods. The total size frequency distribution of the
habitat foraminiferans at the Trevor Channel site was decidedly
skewed to larger-sized individuals, while the cumulative buc-
cal contents were skewed to smaller ones. One explanation
of this shift involves selective and effective removal of most
of the smaller prey by the predators. Similar patterns were
also evident at the other stations but were based on smaller
sample sizes; both the foraminiferan buccal contents, and
habitat foraminiferans were less abundant at the Mayne Bay
and Sandford Island sites (Figs. 16, 19, and 23).

Likewise, the effects of species-specific predation by
the scaphopod predators appeared evident. Typically, the
species that were most common in the habitat were not as
commonly represented in the diets. With the case of Florilus
basispinatus and the Buliminella spp., this shift was particular-
ly evident. The converse was notable with respect to
Cribrononion lene. This species was the most abundant prey

fe ee a
5@) =|
|
40/- | +Cadulus _
i | '-Dentalium|
s [ | —«Pulsellum
a [ |
c Sal |
0 = —
5 ft | |
a r
4 20;
0 i: |
0 | |
5 || F | |
o Kt | i:
||
18} || ! |
a eee |
Mi:eerea © OS eee ll
Lane il |
r | ik ee eee oa oe oes. Pooks Deere sear e 14
ee it ;
—— ooo S| (A es scree
24 32 40 48 56 64 ie
Hours

Fig. 24. The buccal pouch clearing times for all the scaphopod
species; mean values + 1 standard deviation are plotted. For clari-
ty, the standard deviation bars are placed to the left of the times for
Dentalium rectius, and to the right of the times for Pulsellum

salishorum.

SHIMEK: SCAPHOPOD DIETS 167

eaten (Appendix Tables 1-3), but was relatively uncommon
in the habitats (Table 8). While it is tempting to attribute
observed differences to scaphopod predation, and although
| believe this to be the case, such a statement is premature
prior to substantiation based upon experimental data.

ACKNOWLEDGMENTS

This work was supported in large part by the Bamfield Marine
Station. | thank its former director, Dr. Ronald Foreman, for his friendly
support and encouragement of this project. Additional and much wel-
comed financial support for data analysis was provided by the Pacific
Northwest Shell Club. | thank Dr. Donald A. Thomson of the Univer-
sity of Arizona for providing necessary laboratory space and facilities
during a portion of the data analysis. Parametrix, Inc. provided com-
puter support.

Field assistance was provided by Anne Bergey, Janean Clark,
James Dalby, Joel Elliott, Roxie Fredrickson, Tracy Geernaert, Sabina
Leader, Armand Leroi, Steve Rumrill, Ross Warren, and, especially,
the late Sigurd Tveit. | thank R. Fredrickson and Alan J. Kohn for
thorough and rigorous criticisms of earlier drafts of this paper.

| dedicate this paper to Sigurd Tveit, a true Master of Ships,
whose knowledge, skillful help, and friendship made difficult sampl-
ing a pleasure.

LITERATURE CITED

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17:126-138.

Carter, M. S. 1983. Interrelation of shell form, soft part anatomy and
ecology in the siphonodentalioida (Mollusca, Scaphopoda) of
the North West Atlantic continental shelf and slope. Doctoral
dissertation. University of Delaware. 214 pp.

Cockbain, A. E. 1963. Distribution of foraminifera in Juan de Fuca
and Georgia Straits, British Columbia, Canada. Contributions
of the Cushman Foundation for Foram Research 14:37-57.

Dale, N. G. 1974. Bacteria in intertidal sediments — factors related
to their distribution. Limnology and Oceanography 19:509-518.

Davis, J. D. 1968. A note on the behavior of the scaphopod, Cadulus
quadridentatus (Dall) 1881. Proceedings of the Malacological
Society of London 38:135-138.

Dinamani, P. 1964. Feeding in Dentalium conspicuum. Proceedings
of the Malacological Society of London 36:1-5.

Fisher-Piette, E. and A. Franc. 1968. Classe des Scaphopodes. /n:
Grasse, P. P. ed. pp. 987-1017. Traite de Zoologie: Anatomie,
Systematique, Biologie. Mollusques, Gasteropodes et
Scaphopodes. Tome 5: Fasc. Ill.

Gainey, Jr., L. F. 1972. The use of the foot and the captacula in the
feeding in Dentalium. Veliger 15:29-34.

Gallagher, M. T. 1979. Substrate controlled biofacies: Recent
foraminifera from the continental shelf and slope of Vancouver
Island, British Columbia. Doctoral dissertation. University of
Calgary, Canada. 206 pp.

Kozloff, E. N. 1987. Marine Invertebrates of the Pacific Northwest.
University of Washington Press, Seattle. 511 pp.

Lacaze-Duthiers, F. J. H. 1856. Histoire de l’organisation et du
développement du Dentale. Annales des Sciénces Naturelles
(Zool) 4(6):225-281.

Lacaze-Duthiers, F. J. H. 1857. Histoire de l’organisation et du
développement du Dentale. Annales des Sciénces Naturelles
(Zool) 4(7):319-385.

Lankford, R. R. and F. B. Phleger. 1973. Foraminifera from the near-
shore turbulent zone, Western North America. Journal of
Foraminiferan Research 3:101-132.

McFadien, M. S. 1973. Zoogeography and ecology of seven species
of Panamic-Pacific Scaphopods. Veliger 15:340-347.
Meadows, P. S. 1964. Experiments on substrate selection by Cor-
ophium species: films and bacteria on sand particles. Jour-

nal of Experimental Biology 41:499-511.

Morton, J. E. 1959. The habits and feeding organs of Dentalium en-
talis. Journal of the Marine Biological Association of the United
Kingdom 38:225-238.

Pilsbry, H. A. and B. Sharp. 1897-1898. Class Scaphopoda. /n: G. W.
Tyron, Jr. and H. A. Pilsbry, Manual of conchology; Ser. 1, Vol.
17: pp. xxxii+144 (1897); pp. 145-280 (1898), pls. 1-39.

Poole, R. W. 1974. An Introduction to Quantitative Ecology. McGraw-
Hill Book Company. New York. 532 pp.

Poon, Perry A. 1987. The diet and feeding behavior of Caudulus tolmiei
Dall, 1897 (Scaphopoda: Siphonodentalioida). Nautilus
101:88-92.

Shimek, R. L. 1988. The functional morphology of scaphopod cap-
tacula. Veliger 30:213-221.

Shimek, R. L. 1989. Shell morphometrics and systematics: A revi-
sion of the slender, shallow-water Cadulus of the Northeastern
Pacific (Scaphopoda: Gadilida). Veliger 32:233-246.

Sokal, R. R. and F. J. Rohlf. 1981. Biometry. W. H. Freeman and Com-
pany. New York. 859 pp.

STSC, Inc. 1986, 1987, 1988. STATGRAPHICS. Statistical Graphics
Corporation. Rockville, Maryland.

Taib, N. T. 1980. Some observations on living animals of Dentalium
entalis L. Journal of the College of Science, University of
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291 pp.

Date of manuscript acceptance: 8 November 1989.

168

Appendix Table 1. Gut contents of 87 Cadulus aberrans [86 (=98.9%)

AMER. MALAC. BULL. 7(2) (1990)

Appendix Table 2. Gut contenis of 231 Dentalium rectius [144

(=62.3%) feeding].

feeding].
Buccal Contents Total Percent
Item Cumulative

Cribrononion lene 697 27.76 27.76
Elphidiella hannai 373 14.85 42.61
Foraminiferan fragments 339 13.50 56.11
Florilus basispinatus 336 13.38 69.49
Rosalina columbiensis 179 7.13 76.62
Buliminella exilis 99 3.94 80.57
R. columbiensis (test) 69 2.75 83.31
B. elegantissima 46 1.83 85.15
C. lene (test) 42 1.67 86.82
Mineral grains 31 1.23 88.05
B. exilis (test) 28 112 89.17
Nonionella stella 27 1.08 90.24
F. basispinatus (test) 26 1.04 91.28
Foraminiferan sp. 26 1.04 92.31
Textularia sp. 25 1.00 93.31
B. elegantissima (test) 24 0.96 94.27
Globobulimina sp. (test) 17 0.68 94.94
Triloculina sp. 17 0.68 95.62
Uvigerina sp. 16 0.64 96.26
B. sp.C 13 0.52 96.77
E. hannai (test) 11 0.44 97.21
Diatom frustrules 9 0.36 97.57
T sp. B Ff 0.28 97.85
Foraminiferan sp. (test) 6 0.24 98.09
Textularia sp. (test) 6 0.24 98.33
Faujacina sp. 5 0.20 98.53
Nonion sp. D 5 0.20 98.73
Triloculina sp. C 5 0.20 98.92
Rotorbinella sp. 4 0.16 99.08
Astrorhizidae sp. 3 0.12 99.20
Nonionella stella (test) 3 0.12 99.32
Cibicides sp. 2 0.08 99.40
Globobulimina sp. 2 0.08 99.48
Black spherules 1 0.04 99.52
B. sp. C (test) 1 0.04 99.56
B. sp. D (test) 1 0.04 99.60
Discorbinella sp. 1 0.04 99.64
Haplophragmoides sp. 1 0.04 99.68
Lagena sp. D 1 0.04 99.72
Ostracod valve 1 0.04 99.76
Rheophax sp. 1 0.04 99.80
Sacculina sp. 1 0.04 99.84
Sediment bolus 1 0.04 99.88
Spirulina sp. 1 0.04 99.92
Uvigerina sp. (test) 1 0.04 99.96
Virulina sp. 1 0.04 100.00

TOTAL = 47 TAXA

2511

Buccal Contents Total Percent
Item Cumulative

Cribrononion lene 132 20.18 20.18
Sediment bolus 96 14.68 34.86
Mite eggs 64 9.79 44.65
Fecal pellet 42 6.42 51.07
Foraminiferan fragment A 6.27 57.34
Florilus basispinatus 38 5.81 63.15
Mineral grains 37 5.66 68.81
Rosalina columbiensis (test) 22 3.36 72.17
Buliminella elegantissima 15 2.29 74.46
R. columbiensis 15 2.29 76.76
Cribrononion lene (test) 14 2.14 78.90
F. basispinatus (test) 14 2.14 81.04
Arthropod cuticle 11 1.68 82.72
Elphidiella hannai 11 1.68 84.40
Black spherules 7 1.07 85.47
Kinorhynch sp. 7 1.07 86.54
B. elegantissima (test) 6 0.92 87.46
Globobulimina sp. 6 0.92 88.38
B. exilis 5 0.76 89.14
Foraminiferan sp. 5 0.76 89.91
Ostracod valve 5 0.76 90.67
Rhizzamidae sp. 5 0.76 91.44
Astrorhizidae sp. 4 0.61 92.05
Nonionella stella 4 0.61 92.66
Unidentified turbellarian 4 0.61 93.27
Bivalve sp. 3 0.46 93.73
Bivalve valve 3 0.46 94.19
Clear eggs 3 0.46 94.65
Uvigerina sp. 3 0.46 95.11
Brisaster latifrons ossicle 2 0.31 95.41
Flintia sp. 2 0.31 95.72
Globobulimina sp. test 2 0.31 96.02
Haplophragmoides sp. 2 0.31 96.33
Ostracod sp. 2 0.31 96.64
Unidentified planulae 2 0.31 96.94
Sacculina sp. 2 0.31 97.25
Annulated object 1 0.15 97.40
Barnacle cyprid 1 0.15 97.55
Bryozoan statoblast 1 0.15 97.71
Buliminella exilis (test) 1 0.15 97.86
Cibicides sp. 1 0.15 98.01
E. hannai (test) 1 0.15 98.17
Gastropod eggs 1 0.15 98.32
Unidentified mite 1 0.15 98.47
Mollusk shell 1 0.15 98.62
Nonion sp. 1 0.15 98.78
Pegiidae sp. 1 0.15 98.93
Plastic rope 1 0.15 99.08
Polychaete setae 1 0.15 99.24
Textularia sp. 1 0.15 99.39
Triloculina sp. B 1 0.15 99.54
T sp. C 1 0.15 99.69
T sp. A 1 0.15 99.85
Foraminiferan test 1 0.15 100.00

TOTAL = 34 TAXA

654

SHIMEK: SCAPHOPOD DIETS 169

Appendix Table 3. Buccal contents of 149 Pulsellum salishorum [102
(=68.4%) feeding].

Buccal Contents Total Percent
Item Cumulative

Cribonion lene 210 34.83 34.83
Foraminiferan fragment 113 18.74 53.57
Foraminiferan test 45 7.46 61.03
Unidentified foraminiferan 37 6.14 67.16
Mineral grains 35 5.80 72.97
Elphidiella hannai 25 4.15 77M
Rosalina columbiensis 21 3.48 80.60
Florilus basispinatus 18 2.99 83.58
C. lene (test) 14 2.32 85.90
R. columbiensis (test) 12 1.99 87.89
Buliminella exilis 10 1.66 89.55
Sediment bolus 7 1.16 90.71
Nonionella stella 6 1.00 91.71
Sacculina sp. 6 1.00 92.70
Black spherules 5 0.83 93.53
Triloculina sp. A 5 0.83 94.36
B. elegantissima 4 0.66 95.02
Rotorbinella sp. 4 0.66 95.69
Uvigerina sp. 4 0.66 96.35
B. sp. C 3 0.50 96.85
Fecal pellet 3 0.50 97.35
Mite eggs 3 0.50 97.84
B. elegantissima (test) 2 0.33 98..18
B. exilis (test) 2 0.33 98.51
F. basispinatus test 2 0.33 98.84
N. stella 2 0.33 99.17
Dsicorbinella sp. 1 0.17 99.34
Faujacina sp. 1 0.17 99.50
Pegiidae sp. 1 0.17 99.67
Rotorbinella sp. (test) 1 0.17 99.83
Textularia sp. 1 0.17 100.00
TOTAL = 31 TAXA 603

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.00 135.00
30.00 537.00
.00 22.50
26.00 1,093.39
22.00 730.00
550.00 1,972.00
685.50 15,247.89
.00 1,582.89

.00 938.00

.00 393.00

00 1,606.25

.00 135.25

.00 4,655.76

171

$67,581.51

$67,581.51

Other Sales Receipts

Endowment Fund Donations
Interest on All Accounts
Miscellaneous Donations
AMU Registration/Meeting
Separates

Total Other Sales Receipts

Total Cash Receipts

DISBURSEMENTS

Bulletin/Expenses
Bulletin/Postage
Bulletin/Printing

AMU Newsletter/Postage
AMU Newsletter/Printing
AMU Newsletter/Expenses
Other Postage

Other Printing

Office Supplies

Dues

Officer’s Travel
Accounting

Filing Fee (California)

Symposium Endowment Fund Dep.

Student Awards
Insurance

Bank Charges
Miscellaneous/Petty Cash
AMU Meeting

Total Disbursements

Net Income (Loss)

172

00 5,078.65

3,835.00 4,853.60

00 342.39

00 1,990.79

00 279.23

3,835.00 12,544.66

$ 4,520.50 $32,448.31
00

00 1,286.09

00 19,031.36

00 213.21

00 4,212.37

00 351.37

00 500.75

00 112.65

00 417.07

00 610.00

00 2,000.00

00 300.00

00 32.50

00 2,100.00

00 750.00

00 199.00

00 32.50

00 882.22

00 532.00

00 $33,562.91

$ 4,520.50 ($ 1,114.60)

56th ANNUAL MEETING
THE AMERICAN MALACOLOGICAL UNION
WOODS HOLE, MASSACHUSETTS
JUNE 3 - 7, 1990

The 56th annual meeting of the AMU will be held from 3-7 June 1990 at the Marine Biological
Laboratory (MBL) in the village of Woods Hole, Massachusetts on Cape Cod. The MBL recently
celebrated its Centennial and has a distinguished history of research on molluscs. Its library (to which
all registrants will have free access) is considered one of the best in the world. Woods Hole is also
the home of the Woods Hole Oceanographic Institution (WHOIl), National Marine Fisheries Service
(NMFS), U.S. Geological Survey and Sea Education Associates.

Woods Hole is accessible by excellent bus service or by car from Boston or Providence, R.!. (each
70 miles); the nearest major airport is in Boston. Discounted air transportation can be coordinated,
free of charge, through Rhodes Travel (1-800-356-6008). Dormitories (for students only) and motel
accommodations are available in Woods Hole, and a very reasonable cafeteria meal plan will be
available on campus during the meeting.

Two symposia are planned:

THE BEHAVIOR OF MOLLUSCS
With special session on Integrative Neurobiology and Behavior, round-table
discussion on evolutionary aspects of behavior, and a film festival
(Organized by Dr. Roger T. Hanlon)

SYSTEMATICS, BIOLOGY AND FISHERIES OF RECENT CEPHALOPODS
in honor of the late Professor Gilbert L. Voss
(Organized by Dr. Clyde F. E. Roper)

In addition to the symposia, contributed papers and poster presentations, scheduled events will in-
clude a workshop on home aquaria, a marine collecting trip on the MBL vessel in Vineyard Sound,
shore trips to collect and observe intertidal molluscs, trips to the NMFS Aquarium, WHOI, Boston
Aquarium and the National Seashore, plus an auction, outdoor clam bake and a banquet.

Vacation opportunities abound throughout Cape Cod. The ferry to Martha’s Vineyard and Nantucket
is located in Woods Hole. Weather in early June is likely to be quite cool.

For further information please contact:

Roger T. Hanlon
President, AMU
The Marine Biomedical Institute
The University of Texas Medical Branch
Galveston, Texas 77550-2772

Telephone (409) 761-2133
FAX (409) 762-9382

173

AMERICAN MUSEUM OF
NATURAL HISTORY FELLOWSHIPS

FELLOWSHIPS - American Museum of Natural History Research
/Museum Fellowships are available to postdoctoral researchers and
established scholars starting in summer and fall 1990. Deadline for ap-
plications is January 15, 1990.

GRANTS - Grants are available to advanced predoctoral candidates and
recent postdoctoral researchers. Awards range from $200 - $1,000.
Deadlines vary according to grant program:

* Theodore Roosevelt (N.A. fauna) - February 15, 1990

* Lerner-Gray (marine) - March 15, 1990.

Request information booklet and applications from the Office of Grants
and Fellowships, Department |, American Museum of Natural History,
Central Park West at 79th Street, New York, New York 10024, U.S.A.

EXOTIC BIVALVE SYMPOSIUM

In conjunction with the Annual Meeting of the American Society of Lim-
nology and Oceanography (ASLO) to be held in Williamsburg, Virginia,
U.S.A., from 10-14 June 1990, a special symposium examining the distribu-
tion, physiology, and ecosystem role of the Asiatic clam, Corbicula
fluminea, and European zebra mussel, Dreisenna polymorpha, will be held.
Approximately 20 papers will be presented. Other special sessions or sym-
posia at the ASLO meeting will include Eutrophication of Estuaries, Spring
Phytoplankton Blooms, Global Climate Change, Turbidity Dynamics in
Freshwater and Marine Systems, and Extracellular Enzyme Activities in
Aquatic Ecosystems. For further information on the Exotic Bivalve Sym-
posium, contact Thomas Crisman/Robert Brock (904-392-0838) or Renata
Claudi (416-592-4163).

174

INDEX TO VOLUME 7 (1 and 2)

AUTHOR INDEX

Adamkewicz, S. L. 117 Kilgour, B. W. 109 Reynolds, P. D. 137
Aronson, R. B. 47 Liu, Y. Y. 131 Shimek, R. L. 147
Bullock, R. C. 13 Lynn, D. H. 109 Staub, K. C. 93
Counts, C. L., Ill 81 Lyons, W. G. 57 Tan Tiu, A. 65
Davis, G. M. 131 Mackie, G. L. 109 Theler, J. L. 127
Elek, J. A. 117 Morton, B. 73 Upatham, E. S. 93
Hanlon, R. T. 21 Nakamura, H. K. 105 Viyanant, V. 93
Hillis, D. M. 7 Ojima, Y. 105 Wolterding, M. R. 21
Houbrick, R. S. 1 Prezant, R. S. 65 Woodruff, D. S. 93
PRIMARY MOLLUSCAN TAXA INDEX

Acanthopleura 13 Corbiculidae 81 Octopus 21, 47
Amblema 128 Crassinella 57 Paragonimus 131
Anodonta 128 Crassatellidae 57 Psidium 109
Argopecten 117 Dentalium 137, 147 Polymesoda 78
Batissa 73 Fusconaia 128 Potamilis 128
Bulimulidae 7 Halewisia 132 Pulsellum 147
Cadulus 147 Lampsilis 128 Quaadrula 128
Campanile 1 Lasmigona 128 Scaphopoda 137, 147
Campaniloidea 6 Legumia 128 Semisulcospira 105
Cerithoidea 1 Leptodea 128 Tricula 132
Chitonidae 13 Liguus 7 Tritogonia 128
Corbicula 65, 81 Neotricula 93, 131

Corbiculacea 81 Octopodidae 21, 47

Dates of Publication

Volume 7(1), April, 1989
Volume 7(2), February, 1990

wo

| with molluscs will be considered for publication.

on paper, SéabieSpaccds ad all pages numbered con-
=a peiively with numbers appearing in the upper right hand
oe each page. Peeve ample margins on all sides.

sda, raniena 20814, U. S.A.
- Text, when appropriate, should be Saeed in sec-

1. Cover page with title, author(s) and ad-

-dress(es), and suggested running title of no more than

50 characters ‘and spaces -

PSs F 2. Abstract (ess. than 5 percent of manuscript

i as ve | length) ©

4 voles 3. Text of Tneneecripi: starting with a brief in-

Bee oaks ae troduction followed by methodology, results, and dis-

eo ~ cussion. ‘Separate sections of text with centered sub-
— in capital letters.

4. Acknowledgments —

_ 5. Literature cited

PO sone ae 6. Figure captions

Sy e > References should be cited within text as follows: Vail

BF (1977) or (Vail, 1977). Dual authorship should be cited as

follows: Yonge and Thompson (1976) or (Yonge and Thomp-

son, 1976). Multiple authors of a single article should be cited
ees as follows: Beattie et a/. (1980) or (Beattie et a/., 1980).

= - that taxon the first time the name appears in the manuscript
_ [e.g. Crassostrea virginica (Gmelin)). ‘This includes non-

-molluscan taxa. The full generic name along with specific
ES. epithet should be written out the first time that taxon is re-
ferred to in each paragraph. The generic name can be ab-
fee breviated in the erat of the paragraph as follows:
_ C. virginica.
ee In the literature cited section of the manuscript refer-
a ences must also be typed double spaced. All authors must be
lly” identified, listed alphabetically and journal titles must be
ee _ unabbreviated. Citations should appear as follows:
: Beattie, J. H., K. K. Chew, and W. K. Hershberger. 1980.
eget Differential survival of selected strains of Pa-
ries ee __ Cific oysters (Crassostrea gigas) during summer
mortality. Proceedings of the National Shell-
~~ fisheries Association 70(2):184-189.

: Seed. R. 1980, Shell growth and form in the Bivalvia.
In: Skeletal Growth of Aquatic Organisms, D. C.

Press, New York.

eee Vail, V, A. 1977. Comparative reproductive anatomy
of 3 viviparid gastropods. pec ule
16(2):519-540.

a important ae reports, and detailed reviews.

All binomens should include the author attributed to.

. a ee = - Rhoads and R. A. Lutz, eds. pp. 23-67. Plenum

CONTRIBUTOR INFORMATION

Yonge, C. M. and T. E. Thompson. 1976. Living
Marine Molluscs. William Collins Sons & Co.,

Ltd., London. 288 pp.
Illustrations should be clearly detailed and readily
reproducible. Maximum page size for illustrative purposes is
17.3 cm x 21.9 cm. A two-column format is used with a

- single column being 8.5 cm wide. All line drawings should be

in black, high quality ink. Photographs must be on glossy,
high contrast paper. All diagrams must be numbered in the
lower right hand corners and adequately labeled with suf-

ficiently large labels to prevent obscurance with reduction by
one half. Magnification bars must appear on the figure, or the

caption must read Horizontal field width = xmm or xm. All
measurements must be in metric units. All illustrations sub-
mitted for publication must be fully cropped, mounted on a
firm white backing ready for reproduction, and have author’s
name, paper title, and figure number on the back. All figures
in plates must be nearly contiguous. Additional figures sub-

_ mitted for review purposes must be of high quality reproduc-
tion. Xerographic reproduction of photomicrographs or any

detailed photographs will not be acceptable for review.
Abbreviations used in figures should occur in the figure
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which figures should appear. Color illustrations can be in-
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be returned to author if requested.

Any manuscript not conforming to AMB format will be
returned to the author.

Proofs. Page proofs will be sent to the author and must be
checked for printer’s errors and returned to the printer within
a three day period. Changes in text other than printer errors
will produce publishing charges that will be billed to the
author.

Charges. There are no mandatory pages costs to authors
lacking financial support. Authors with institutional, grant or
other research support will be billed for page charges. The
current rate is $30.00 per printed page.

Reprints. Order forms and reprint cost information will be

sent with page proofs. The author receiving the order form
is responsible for insuring that orders for any coauthors are
also placed at that time.

Submission. Submit all manuscripts to Dr. Robert S. Prezant,
Editor-in-Chief, American Malacological Bulletin, Department
of Biology, Indiana University of Pennsylvania, Indiana,
Pennsylvania, 15705, U.S.A.

Subscription Costs. Institutional subscriptions are avail-
able at a cost of $28.00 per volume. [Volumes 1 and 2 are
available for $18.00 per volume.] Membership in the Ameri-
can Malacological Union, which includes personal subscrip-
tions to the Bulletin, is available for $20.00 ($15.00 for
students) and a one-time initial fee of $1.50. All prices quoted
are in U.S. funds. Outside the U.S. postal zones, add $3.00

~ seamail and $6.00 airmail per volume or membership. For

subscriptions or membership information contact AMU

Secretary-Treasurer, Dr. Clement L. Counts III, University of

Maryland, Eastern Shore, Box 1106, Princess Anne, MD 21853.

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