Full text of "Journal of shellfish research"

JOURNAL OF SHELLFISH RESEARCH

VOLUME 3, NUMBER 1

JUNE 1983

The Journal of Shellfish Research (formerly Proceedings of the

National Shellfisheries Association) is the official publication

of the National Shellfisheries Association

Editor

Dr. Robert E. Hillman

Battelle

New England Marine Research Laboratory

Duxbury, Massachusetts 02332

Managing Editor

Dr. Edwin W. Cake, Jr.
Gulf Coast Research Laboratory
Ocean Springs, Mississippi 39564

Associate Editors

Dr. Jay D. Andrews

Virginia Institute of Marine Sciences

Gloucester Point, Virginia 23062

Dr. Anthony Calabrese
National Marine Fisheries Service
Milford, Connecticut 06460

Cornell University
Ithaca, New York 14853

Dr. Richard A. Lutz
Nelson Biological Laboratories
Rutgers University
Piscataway, New Jersey 08854

Dr. Kenneth K. Chew
College of Fisheries
University of Washington
Seattle, Washington 98195

Dr. Gilbert Pauley
College of Fisheries
University of Washington
Seattle, Washington 98195

Dr. Paul A. Haefner, Jr.
Rochester Institute of Technology
Rochester, New York 14623

Dr. Daniel B. Quayle
Pacific Biological Laboratory
Nanaimo, British Columbia, Canada

Dr. Herbert Hidu
Ira C. Darling Center
University of Maine
Walpole, Maine 04573

Dr. Louis Leibovitz

New York State College of Veterinary Medicine

Dr. Aaron Rosenfield

National Marine Fisheries Service

Oxford, Maryland 21654

Dr. Frederic M. Serchuk
National Marine Fisheries Service
Woods Hole, Massachusetts 02543

Journal of Shellfish Research

Volume 3, Number 1

ISSN: 00775711

June 1983

Journal of Shellfish Research, Vol. 3, No. 1, 1-9, 1983.

PREDATION OF JUVENILES OF THE HARD CLAM MERCENARIA MERCENARIA (LINNE)

BY THE SNAPPING SHRIMP ALPHEUS HETEROCHAELIS SAY

AND ALPHEUS NORM ANNI KINGSLEY

BRIAN F. BEAL 1

The University of North Carolina at Chapel Hill
Institute of Marine Sciences
Morehead City, North Carolina 28557

OCT 5 1984

ABSTRACT Two species of snapping shrimp, Alpheus heterochaelis and A. normanni, collected near Beaufort, North
Carolina, during June 1982, and then held in the laboratory, used their major chelae to crush and consume juveniles of the
hard clam Mercenaria mercenaria. Snapping shrimp (19.1 to 39.4 mm in total body length (TL] ) ate clams in the largest
size-class (15.1 to 20.0 mm in shell length), but preferred smaller clams when offered equal numbers in this large size-class
and in each of three smaller size-classes. Female snapping shrimp, regardless of species, exhibited a statistically higher
predation rate than males when the results of five separate experiments were combined. The major chelae of the females of
specimens of A. heterochaelis (>32.0 mm TL) were smaller than those of equal size males. Alpheus heterochaelis (19.1 to
27.2 mm TL) had a larger major chela for a given body length than did specimens of A. normanni; however, predation rates
of the two species were not significantly different. The number of clams crushed was related to both the size of the major
chelae and total body length for A. normanni, but not for A heterochaelis. Alpheus spp. inflict two types of shell damage
which are identical to those caused by blue crabs. These results imply that previous studies may have overestimated the
importance of crab predation and underestimated or ignored the importance of predation by snapping shrimp.

KEY WORDS: Alpheus, snapping shrimp, predation, Mercenaria, hard clams

INTRODUCTION

The hard clam or northern quahog Mercenaria mercen-
aria (Linne) is distributed along the Atlantic coast from the
Gulf of St. Lawrence to the northern Gulf of Mexico and
occurs intertidally down to 1 5 m (Menzel 1970). This species
is harvested commercially throughout most of its range;
e.g., during 1981 and 1982 in North Carolina, the hard
clam fishery ranked third in importance of all commercial
fisheries based on a dockside dollar value of $5.4 million
and $6.6 million, respectively (Street 1982).

A progression of predators follows the growth of the hard
clam from the earliest planktonic (Loosanoff 1959, Carriker
1961), post-settlement (Hunt 1981), and juvenile stages
(Carriker 1951, Goodwin 1968, Whetstone and Eversole
1978) through adulthood (Carriker 1951, MacKenzie 1977,
Greene 1978, Peterson 1982). As M. mercenaria increases
in size so does its predators; because large predators are
more commonly recognized in the field and have been
studied extensively in the laboratory, their importance in
regulating hard clam population sizes may have been over-
emphasized. Investigations of predation on natural or
hatchery-reared juvenile hard clams by blue crabs (Callinectes
sapidus Rathbun) (Carriker 1951, Menzel and Sims 1964,
Castagna and Kraeuter 1977), mud crabs (various xanthid
genera) (Landers 1954, MacKenzie 1977, Whetstone and
Eversole 1978), and miscellaneous species (Menzel et al.
1976) imply that those predators are responsible for the

Present address: The University of Maine at Orono, Cooperative
Extension Service, 5 Cooper St., P.O. Box 189, Machias, ME 04654

majority of natural post-settlement mortality of hard clams.
Resource managers and fishermen who operate commercial
bottom leases should be aware of the potential effectiveness
of these predators in reducing hard clam populations.

I conducted a series field experiments near Beaufort, NC,
from August 1981 through April 1982, in which juveniles
of M. mercenaria (6.0 to 1 5.0 mm in length) were maintained
in cages designed to exclude large (> 6.4 mm) epibenthic
predators (Beal, unpublished data). Because numerous
snapping shrimp were observed inside the field cages, which
also contained several crushed juvenile hard clams, they
were suspected of being an important additional consumer
of juvenile clams.

As a result of these field investigations, I performed
several laboratory experiments that clearly showed that two
species of snapping shrimp, Alpheus heterochaelis Say and
Alpheus normanni Kingsley, should be added to the list of
known hard clam predators. Here I demonstrate that both
species will crush and consume juvenile hard clams under
laboratory conditions and provide field observations that
indicate they do so in nature as well. Several factors are
also examined:

1. Is size of snapping shrimp correlated with its preda-
tion rate?

2. Do shrimp show a size preference within the size-
classes of clams they are able to crush?

3. Does sex or species of snapping shrimp affect
predation rate?

4. Can clam mortality, caused by blue crabs, be distin-
guished from that inflicted by snapping shrimp on
the basis of shell damage? ^

BEAL

MATERIALS AND METHODS

Snapping shrimp and shell debris were obtained from
two oyster rocks (reefs) near Beaufort, NC, on 18 and
26 June 1982, using a suction dredge. Shell debris (hash)
was the substrate used in all experiments and consisted of
dead and fragmented oysters and clams greater than 3.0 mm
(the smallest size the dredge efficiently captured). Juvenile
hard clams were purchased from a commercial dealer and
given a color dot (Mark-Tex Corp., paint) on both valves
(near the umbo) which distinguished them from any dead
clams within the shell debris.

Snapping shrimp and shell debris were brought to the
laboratory on the same day they were collected. Shrimp
were placed in glass finger bowls where they were given
crushed hard clams as food. Bowls were placed in large
tanks (75 X 75 X 30 cm) supplied with unfiltered seawater.
No snapping shrimp were held longer than four days in the
pre -experimental setting.

Shell debris was sieved through a 3.2-mm mesh to remove
all fine sediments and small benthos at the beginning of
each experiment. Any large animals were also removed
before the shell debris was placed in finger bowls (20.0 cm
dia;6.5-cm depth) to a depth of 4 cm.

Forty marked clams were placed at a depth of 1 cm
before one snapping shrimp was added to each bowl in each
experiment. Nylon window screening (1.2-mm mesh) was
placed over the top of each bowl and secured by an elastic
band to ensure that the shrimp remained inside the bowl
during the course of the experiment. Controls were
employed to separate all types of shrimp-caused mortality
from all other sources of mortality. The controls were
treated identically to the other clams placed in finger
bowls except they received no snapping shrimp.

Each tank held nine finger bowls and in experiments
where more than one tank was used, treatment and control
bowls were randomly assigned to tanks. The nylon tops
were cleaned daily using hands to brush away accumulated
silt; the bowls were not removed from the tanks. Snapping
shrimp were removed from each bowl and the contents
of the bowls were sieved through 1-mm mesh after one week.
Recovered clams were separated into three categories:
living, dead (empty, undamaged shells), or dead (crushed).

Table 1 shows the experimental interval, the number of
replicate Alpheus spp. used, and the number of controls
for each experiment. Experiments A through C were
designed to test whether A. heterochaelis could crush and
consume juvenile hard clams. The same two snapping shrimp
were used in both experiments A and B. Replication was
increased in experiments D and E because of the large
variability in crushing rates of the snapping shrimp.

The major chela (MC) of each snapping shrimp was
measured from the distal end of the dactylus longitudinally
to the proximal end of the propodus, and total body length
(TL) was measured from the rostrum to the telson after
every experiment. These two morphological traits were

measured to test whether the relationship between size
of the MC and TL differed between sexes of large specimens
of A. heterochaelis and between species of smaller snapping
shrimp. In addition, I tested whether predation rate was
related to either morphological trait.

TABLE 1.

The experimental interval, number of Alpheus, and
number of controls for each experiment.

Number of Alpheus

Number of

Experiment

Dates

spp. treatments

Controls

18 June to :

25 June

2 (A. heterochaelis)

25 June to

2 July

2 (.4. heterochaelis)

26 June to

3 July

4 (A. heterochaelis)

29 June to

6 July

14 (A. heterochaelis)

30 June to

7 July

12 (A. heterochaelis)
8 (A. normanni)

Four size-classes of juveniles of M. mercenaria (6.0 to
8.0, 8.1 to 10.0, 10.1 to 15.0, and 15.1 to 20.0 mm in shell
length [SL, the greatest anterior to posterior measurement] )
were used to test if shrimp preferred clams within a certain
size. Ten clams from each size category were placed in each
bowl. A total of 20 large specimens of A. heterochaelis
(mean TL = 34.1 mm ± 2.5 SD) was used in these experi-
ments. To determine the effects of sex of snapping shrimp
on predation rate, the nonparametric Wilcoxon two-sample
test on total number crushed by individual snapping shrimp
was used. Data from experiments A through D were
combined because ( 1 ) the time interval for each experiment
was identical (7 days); (2) there was no apparent effect of
time on predation rate; and (3) size categories of juvenile
hard clams, as well as number of clams used within each
size category, were held constant. Mean total numbers
crushed by individual shrimp were used from experiments
A and B because the same shrimp were used in both trials.
Total counts were used for individual shrimp in experi-
ments C and D. Morphometric data from experiments A
through D were combined and the lines expressing TL to
MC for the 1 1 male snapping shrimp (Y = 2.99 + 0.487X;
r 2 = 0.74) and 9 female snapping shrimp (Y = 5.03 +
0.323X; r 2 = 0.69) were compared using multiple regression
analysis.

In experiment E, individuals of both species were smaller
than those specimens of A. heterochaelis used in the
previous experiments. Twelve specimens of A. heterochaelis
(mean TL = 23.4 mm ± 2.6 SD) and eight specimens of
A. normanni (mean TL = 24.0 mm ± 1.9 SD) were used.
Clams from only two size-classes (4.5—8.0 mm and 8.1—
10.0 mm) were used because of the small size of these
snapping shrimp. Twenty clams from each size category
were placed in each bowl. A Model I 2-way analysis of
variance (ANOVA) was performed on numbers crushed to
test the effects of species and sex of snapping shrimp on

Predation of Juvenile clams by snapping Shrimp

predation rate. Numbers crushed (Y) were first transformed
with In (Y + 1 ) and a Bartlett's test (Sokal and Rohlf
1969) was performed to determine whether the transforma-
tion helped eliminate variance heterogeneity. Morphometric
data from male snapping shrimp were pooled with data
from female shrimp for each species in experiment E to
determine whether the two species differed in their relation
between TL and MC for the 1 2 specimens of A. heterochaelis
(Y = 2.71 + 0.382X; r 2 = 0.37) and the 8 specimens of
A.normanni{y = 3.55 + 0.493X;r 2 = 0.56). Again, multiple
regression analysis was used to compare lines.

Five specimens of A. heterochaelis were placed in
isopropyl alcohol within 12 hours after feeding to test
whether shell fragments pass through the cardiac stomachs
of snapping shrimp. After one hour the cardiac stomach of
each shrimp was excised and examined.

Temperature and salinity were monitored daily within
each tank. Tanks never differed by more than 0.7°C or
1 ppt S on any given day. The temperature range for the
entire experimental interval (18 June to 7 July) was 24.3 to
27.5°C. The salinity range for the same time interval was
32 to 34 ppt S.

Four blue crabs, Callinectes sapidus Rathbun (carapace
widths: 146.9, 136.7, 74.8 and 59.7 mm), were placed in
separate seawater tanks (25 X 25 X 30 cm) without sedi-
ment but containing 40 juvenile hard clams (10 from each
size category used in experiments A through D) to compare
shell damage inflicted by Alpheus spp. with that described
for crabs (Venneij 1978). The crabs were used to test
whether it is possible to correctly assign clam mortality to
the proper predator on the basis of shell damage. Crabs
remained in the tanks until at least 50% of the clams had
been crushed. This took 3 days for the smallest blue crab
and 3 hours for the largest.

RESULTS

Experiments A through D (Table 2)

No clam mortalities occurred in the control bowls, but
a total of 77 clam deaths occurred in those bowls containing
the snapping shrimp A. heterochaelis; in each case a chipped
or crushed clam shell was recovered. This clearly demon-
strates that snapping shrimp crush juvenile hard clams;
furthermore, body tissues were removed from each valve
indicating that the clams were eaten.

I observed a female of A. heterochaelis (35.2-mm TL)
crush and consume a juvenile hard clam (~ 8.0-mm SL) in a
small finger bowl (10-cm, dia; 5-cm depth) containing no
shell, other substrate, or other clams. The snapping shrimp
grasped the clam near the umbo with the minor chelae, then
lifted the clam several millimeters off the bottom. With
the dactylus cocked, the snapping shrimp raised its major
chela so that the clam was wedged (anterior to posterior
and 2 to 3 mm ventral of the umbo) between the propodus
and dactylus with its umbo and dorsal margin straight up.
The dactylus closed quickly fracturing most of the clam,

leaving only a small portion of the umbo intact. Initially,
the mantle held the fractured pieces of clam together, but
after the shrimp used its minor chela to tear the mantle
from the valve remnants, the small fragments of shell
became separated. The shrimp then tore off pieces of body
tissue and used its minor chela and pereiopods in feeding.
The cardiac stomach of each snapping shrimp examined
contained shell fragments and, in one case, the painted
portion of the clam.

Female snapping shrimp accounted for 92% of all clams
crushed in experiments A through D; however, this was not
statistically significant (P = 0.09). Snapping shrimp showed
a statistical preference for smaller juvenile clams in a chi-
square (X 2 ) test (X 2 = 34.8. df = 3, P < 0.001); 49% of all
the clams crushed and consumed belonged to the smallest
(6.0 to 8.0-mm SL) size-class. Clams were eaten in all
size-classes including the largest (15.1 to 20.0-mm SL).

The variances around the straight lines relating TL to
MC for the 1 1 males and 9 females of A. heterochaelis
(Figure 1) were not significantly different. The lines were
parallel (P > 0.75), but not coincident (P < 0.001 in partial
F-test). Analysis of covariance (ANCOVA) demonstrated
that, even though females had a greater mean TL (35. 41 mm)
than males (32.95 mm), males had a larger MC for a given
TL than females (P < 0.001). Because of the apparent
effect of sex on predation rate in experiments A through D,
sexes were not combined when I tested whether predation
rate could be explained by either morphological trait. No
significant relationships existed between TL (r . = 0.48,
n = ll:r 9 = 0.14, n = 9) or size of MC (r d =0.52, n= 11;
ro = 0.43, n = 9) and predation rate.

Experiment E (Table 3)

One 5.6-mm SL clam died in a control bowl as a result
of natural causes; however, 31 clams died as a result of
crushing in bowls containing A. heterochaelis and 38 clams
were crushed in bowls containing A. normanni. All 31 clams
eaten by specimens of A. heterochaelis in experiment E
belonged to the smaller size-class (4.5-8.0 mm); none were
eaten in the larger size-class (8.1-10.0 mm) as were crushed
and consumed by larger specimens of A. heterochaelis in
experiments A through D. Similarly, 95% of those clams
crushed and consumed by A. normanni came from the
smaller size category. Bartlett's test demonstrated that the
logarithmic transformation reduced variance heterogeneity
and the Model I 2-way ANOVA resulted in no species X sex
interaction (P > 0.50) or effect of species (P > 0.75). The
15 female snapping shrimp ate 67 of the 69 (97%) clams;
the remaining 2 crushed clams were eaten by one of the five
male shrimp. This was not statistically significant (P = 0.065 ).

The straight lines relating TL to MC (Figure 1 ) from
experiment E had equal variances (P > 0.05) and were
parallel (P > 0.75), but not coincident (P < 0.001 in a
partial F-test). Application of ANCOVA yielded a significant
difference (P < 0.001) in the adjusted MC lengths between

BEAL

TABLE 2.

Results of Experiments A through D in which Alpheus heterochaelis was exposed for
7 days to 10 clams in each of four size categories.

Sex

TL*
(mm)

MCf

(mm)

Number Crushed With

in a Size Category (mm)

Total Crushed

Experiment

6.0-8.0

8.1-10.0

10.1-15.0

15.1-20.0

Number Mive

34.8

20.1

38.4

17.6

Control 1

Control 2

34.8

20.1

38.4

17.6

Control 1

Control 2

32.4

18.7

34.0

19.9

34.0

15.9

35.2

16.4

Control 1

Control 2

29.9

17.2

30.0

17.2

30.4

18.0

30.9

19.0

34.4

19.9

34.6

19.9

34.9

18.5

35.9

21.1

32.1

16.2

33.5

15.6

35.0

16.3

35.3

15.4

35.8

16.6

39.4

18.1

Control 1

Control 2

Control 3

Total number

of controls

360

Total number

of males

474

Total number

of females

329

*TL = total body length
fMC = length of major chela

species. Alpheus heterochaelis in the size range 19.1 to
27.2 mm had a larger MC for a given TL than A. normanni
There was no significant (P > 0.05) relationship between
either TL (r = -0.24, n = 12) or length of MC (r = -0.05,
n = 12) and number of clams crushed by A. heterochaelis;
however, predation rate was significantly (P < 0.05) corre-
lated for TL (r = 0.76, n = 8) and MC size (r = 0.77, n = 8)
for A normanni.

Effect of Sex on Predation Rate

Fischer's technique of combining probabilities from
independent tests of significance (Sokal and Rohlf 1969)
was applied to test the effect of sex of snapping shrimp on
predation rate from all experiments. This test resulted in a

significant (P = 0.04) overall effect of sex implying that
females had a greater crushing rate over all experiments.
The effect of sex in experiment E included information
from both species; however, because there was no species X
sex interaction, this test was justified over all experiments.
The size distributions of males and females used in all
experiments were compared because size of snapping shrimp
may influence predation rate. Size of snapping shrimp was
statistically independent of sex (X 2 = 9.38;df =6;P=0.195)
over all experiments.

Shell Damage Inflicted by Snapping Shrimp (Figure 2) and Blue Crabs

Two types of shell damage caused by snapping shrimp
were distinguished by visual inspection. In the first (Type I)

Predation of Juvenile Clams by Snapping Shrimp

E
E

° a A. normanni
t <■ A. heterochaelis

25 30

Total length (in mm)

Figure 1. Relationships between total body length (TL) and size of major chela (MC) for snapping shrimp used in all experiments. Capital
letters refer to experiment. Open circles: Alpheus normanni; closed circles: Alpheus heterochaelis.

at least one of the valves remained intact. Shell chips or
fractures were restricted along the posterior edge and often
both valves had symmetrical chips. Where both valves were
not chipped identically, one valve was chipped along the
posterior edge while damage to the other valve ranged from
restricted ventral margin fractures to an extensively broken
valve having only the umbo region intact. Valves exhibiting
damage of the second type (Type II) had been completely
crushed and only the immediate area around the umbo
was left intact and held together by the hinge ligament
(Figure 2).

To learn if shell damage inflicted by snapping shrimp
and blue crabs was distinguishable, crushed shells from
experiments A through D and from the blue crab experi-
ment were collected and separated by size-class into damage
types. Both predators caused Type I and Type II damage in
each size-class. Sixty clams were crushed by A. hetero-
chaelis in the size range 6.0 to 10.0 mm; 92% exhibited
Type II damage, whereas 53% of the crushed clams between
10.1 and 20.0 mm exhibited Type I damage. Type II damage
occurred in 70% of the juvenile hard clams (6.0 to 10.0 mm
SL) crushed by blue crabs, whereas 8% of the crushed clams
between 10.1 and 20.0 mm suffered Type I damage from
blue crabs.

DISCUSSION

Experiments A through E demonstrate that two species
of snapping shrimp, A. heterochaelis and A. normanni, can
crush and consume juveniles of M. mercenaria and can also
discriminate between sizes of prey when offered a choice.
It is sometimes difficult to relate laboratory experiments
to field experiments because the number of variables
permitted to vary in each is different (Dayton and Oliver
1981); however, two observations from my caging study in
the field suggested that snapping shrimp do indeed prey on
juvenile hard clams (6.0 to 15.0 mm SL) in nature. Snapping
shrimp were found inside complete 1 m 2 cages (6.4-mm
mesh; see Beal [1983] for a detailed cage description)
designed to keep large, epibenthic predators from preying
on juvenile hard clams (Beal, unpublished data). When the
contents of these cages were sieved in November 1981 and
in April 1982, I found live clams as well as shell fragments
which were identical in appearance to those clams crushed
and consumed by Alpheus spp. in this study. No other
predators or signs of predators were observed inside
complete cages.

Female snapping shrimp exhibited a higher predation
rate than did males over all experiments; however, the
mechanism for this behavior was not investigated. Elner and

BEAL

TABLE 3.

Results of Experiment E in which Alpheus heterochaelis and Alpheus normanni
were exposed for 7 days to 20 clams in each of two size categories.

Species

Sex

TL*

(mm)

MCf

(mm)

Number Crushed Within a Size Category (mm)

4.5-8.0

8.1-10.0

Total Crushed

Number Alive

Alpheus heterochaelis

21.4

10.0

25.4

13.7

26.9

12.4

Total males

19.1

10.7

20.3

9.1

20.8

11.2

22.9

12.7

23.2

12.5

23.5

11.5

24.2

9.5

25.6

14.2

26.9

12.1

Total females

Total A. heterochaelis

Alpheus normanni

22.5

7.2

23.4

7.6

Total males

21.6

8.1

23.1

7.8

23.4

7.6

23.8

8.3

26.1

8.1

27.2

11.1

Total females

Total A. normanni

Control 1

Control 2

Control 3

*TL = total body length

f MC = length of major chela

2
2

118

331

449

202

282

Hughes (1978) examined the diet of the shore crab Carcinus
maenus (Linnaeus) and, to avoid potential biases caused by
sexual differences in morphology and predatory behavior,
used only male crabs. Here both sexes were used and, at
least for larger specimens of Alpheus heterochaelis, females
had a smaller major chela than did males of a similar body
length. Because the major chela is used in crushing juvenile
hard clams, males should have had the highest predatory
rate. Ennis (1973) found a difference in the feeding activity
between sexes of the American lobster Homants americanus
Milne Edwards; females continued to feed at a higher level
longer into the winter than did males. Ennis (1973)
suggested that this may have been caused by greater physio-
logical demands on the female due to gonadal development.
If an energetic explanation were true for snapping shrimp,
similar experiments using females with developing versus
developed gonads or, perhaps, immature (juvenile) versus
mature females as well as males would be needed.

Accounts of snapping shrimp as predators are rare.
Hazlett (1962) determined that a species of Alpheus from
Bermuda was omnivorous. Goldberg (1971) studied a species
of Synalpheus in the Florida Keys which preyed upon the
gastropod Coralliophila caribaea Abbott without crushing
it. The shrimp lifted the flexible operculum with its major
chela exposing the gastropod while the minor chela tore
off pieces of the foot. I am unaware of any account of
predation by either A. heterochaelis or A. normanni on a
bivalve mollusc.

Previous investigations concerning the role that the major
chela plays in the behavior and ecology of these snapping
shrimp suggest that it is used agonistically during intra-
and interspecific interactions (Nolan and Salmon 1970,
Schein 1977). Conover and Miller (1978) described the
importance of the major chela in determining the success
of a shrimp in competing for shelter. Glynn (1976) described
a species of snapping shrimp off the Pacific coast of Panama

Predation of Juvenile Clams by snapping shrimp

Figure 2. The size range of the five size-classes of juvenile hard clams and the shell damage caused by Alpheus heterochaelis from experi-
ment A through E. Damage in the smaller size-classes was similar for both species. Each tick mark represents 1 mm.

BEAL

which repulsed the crown-of-thorns sea star and prevented
it from preying on a branching coral. In this study the
major chela of A. heterochaelis (29.9 to 39.4 mm TL) was
smaller in females compared with equal size males. Nolan
and Salmon (1970) noted this sexual dimorphism in both
species. They showed that when a female approached a
larger male, she was threatened and quickly retreated
because of aggressive male snapping; if the TL of a female
was greater than that of the male she approached, the
encounter would continue until cues important in sexual
discrimination could be exchanged.

Whetstone and Eversole( 1978) investigated the predators
of juvenile hard clams in a South Carolina sound. They
collected 13 species of crustaceans from sub tidal and
intertidal trays containing juvenile hard clams over a 19-
month interval and examined their gut contents. They
concluded, on the basis of shell fragments in the cardiac
stomachs (as well as overall numbers collected), that the
xanthid crab Panopeus herbstii Milne Edwards (1,465
collected from May 1975 through December 1976) was the
most important predator of juvenile hard clams. Alpheus
heterochaelis was the second most abundant crustacean
found by Whetstone and Eversole ( 1 84 collected during that
same time interval); nine specimens of A. normanni were
also collected during that study. Whetstone and Eversole
(1978) found no shell fragments in either species of Alpheus
they examined and, on this basis, concluded that snapping
shrimp were not hard clam predators; however, shell frag-
ments were found in the cardiac stomachs of every snapping
shrimp I examined. There may be several reasons why shell
fragments were found in the cardiac stomachs of the
snapping shrimp from this study and not in Whetstone and
Eversole's (1978) investigation:

1. The snapping shrimp they collected may not have
crushed any juvenile hard clams; Whetstone and
Eversole (1978) used hard clams with a mean SL
of 13 mm (however, 19% of the hard clams con-
sumed in my experiments A through D were 10.1
to 15.0 mm SL [Table!]);

2. The snapping shrimp may have been collected or
preserved after evacuation of the cardiac stomachs
had occurred; or

3. The shell fragments may have dissolved in the 10%
formalin solution they used as a preservative.

The results presented in this paper suggest that Alpheus spp.
may be an important predator of juveniles (< 20.0 mm SL)
of M. mercenaria in South Carolina sounds.

I have seen or heard snapping shrimp in a variety of areas
in Bogue, Back, and Core sounds in North Carolina. These
areas have several aspects in common. They either have
muddy substrates with natural shelters such as living or dead
oysters, or seagrass beds. Nolan and Salmon (1970) collected
both species near Beaufort among clumps of oyster shells,
as well as in eelgrass beds. Alpheus heterochaelis was more

often found in muddy areas associated with clumps of
oysters; A. normanni was found primarily in eelgrass beds.
Hoff Stuart (National Marine Fisheries Service, Beaufort,
NC, pers. comm.) found a mean of 6.1 adults of A. normanni
and 1.1 adults of A heterochaelis (TL > 20.0 mm) per m 2
in a Back Sound eelgrass bed during 1975—1976. The mean
number of clams consumed per snapping shrimp per day in
my laboratory experiments was 0.72. This figure is indica-
tive of clams < 15.0 mm SL because only two clams were
consumed that were > 15.0 mm SL. Thus, if that rate is
representative of their hard clam predation in nature,
snapping shrimp of this size in that eelgrass bed may con-
sume approximately 125 clams (4.5 to 15.0 mm SL) per
m 2 per month.

The type of shell damage inflicted by these snapping
shrimp is typical of crabs (Vermeij 1978). Cake (1970)
found that C. sapidus could open large specimens of the
sunray venus clam Macrocallista nimbosa (Lightfoot)
without breaking their shells by "inserting the finger and
cutting the adductor muscles." That type of shell damage
by Callinectes, which leaves behind minute scars of cheliped
activity on the periostracum, was not observed in this study;
in fact, both snapping shrimp and blue crabs inflict similar
types of shell damage. The entire clam is either broken into
bits leaving only the umbo region, or is marginally damaged
with chips occurring around the posterior edge of at least
one valve. According to the results of this study, past
investigations in which clam mortalities were assigned a
particular crushing predator based on shell damage may
have overestimated the importance of crab predation and
underestimated or ignored the importance of predation by
snapping shrimp. Furthermore, commercial clam cultunsts
need to be concerned about protecting seed clams from
snapping shrimp as well as from crabs and other predators.
The spatial distribution and abundance of the bottom-
dwelling snapping shrimp, as well as their natural predation
rates on small hard clams, must be determined to fully
assess the importance of these findings.

ACKNOWLEDGMENTS

I am indebted to G. W. Safrit, Jr., who provided many
of the snapping shrimp used in the laboratory experiments.
K. Bowers, M. E. Colby and S. Smith assisted in the field
and laboratory. R. J. Beal also helped in the laboratory.
H. J. Porter aided in describing types of shell damage.
H. E. Page took the photographs and V. Page prepared the
figure. H. Stuart supplied density data from his dissertation
work. F. J. Schwartz provided computer funds and S. R.
Fegley dissected the snapping shrimp. Additionally, I thank
W. G. Ambrose, Jr., D. R. Colby, P. B. Duncan. S. R.
Fegley, C. H. Peterson, M. C. Watzin, and an anonymous
reviewer for helpful suggestions on earlier drafts of this
manuscript.

Predation of Juvenile Clams by Snapping Shrimp

D. R. Colby, P. B. Duncan, S. R. Fegley, and C. H.
Peterson assisted with experimental design, statistical
analyses, and writing.

Financial support was provided by the Curriculum in
Marine Sciences. University of North Carolina, Chapel

Hill, NC, and the Institute of Marine Sciences, Morehead
City, NC. Support was also provided by the Office of Sea
Grant, NOAA, U.S. Department of Commerce under
Grant No. NA81AA-D-0026, North Carolina Depart-
ment of Administration to C. H. Peterson.

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Beal, B. F. 1983. Effects of environment, intraspecific density,
predation by snapping shrimp and other consumers on the popu-
lation biology of Mercenaria mercenaria near Beaufort, North
Carolina. Chapel Hill, NC: Univ. of North Carolina. 181 p. Thesis.

Cake, E. W., Jr. 1970. Some predator-prey relationships involving
the sunray venus clam, Macrocallista nimbosa (Lightfoot)
(Pelecypoda: Veneridae), along the Gulf coast of Florida.
Tallahassee. FL: Florida State Univ. 166 p. Thesis.

Carriker. M. R. 1951. Observations on the penetration of tightly
closing bivalves by Busycon and other predators. Ecology
32:73-83.

. 1961. Interrelation of functional morphology, behavior,

and autecology in the early stages of the bivalve Mercenaria
mercenaria. J. Elisha Mitchell Sci. Soc. 77:168-241.

Castagna, M. & J. N. Kraeuter. 1977. Mercenaria culture using
stone aggregate for predator protection. Proc. Natl. Shellfish.
Assoc. 67:1-6.

Conover, M. R. & D. E. Miller. 1978. The importance of the large
chela in the territorial and pairing behavior of the snapping
shrimp, Alpheus heterochaelis. Mar. Behav. Physiol. 5:185-192.

Dayton, P. K. & J. S. Oliver. 1981. An evaluation of experimental
analyses of population and community patterns in benthic
marine environments. Tenore, K. R. and B. C. Coull. eds. Marine
Benthic Dynamics. Columbia, SC: Univ. of South Carolina
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Elner, R. W. & R. N. Hughes. 1978. Energy maximization in the
diet of the shore crab, Carcinus maenas. J. Anim. Ecol. 47 :
103-116.

Ennis, G. P. 1973. Food, feeding, and condition of lobsters. Homarus
americanus. throughout the seasonal cycle in Bonavista Bay.
Newfoundland./ Fish. Res. Board Can. 30:1905-1909.

Glynn, P. W. 1976. Some physical and biological determinants of
coral community structure in the Eastern Pacific. Ecol. Monogr.
46:431-456.

Goldberg, W. M. 1971. A note on the feeding behavior of the
snapping shrimp Synalpheus fritzmuelleri Coutiere (Decapod:
Alpheidae). Crustaceana (Leiden) 21:318-320.

Goodwin, W. F. 1968. The growth and survival of planted clams,
Mercenaria mercenaria, on the Georgia Coast. Ga. Game Fish
Comm.. Mar. Fish. Div., Contrib. Ser. No. 9:1-16.

Greene, G. T. 1978. Population structure, growth and mortality of

hard clams at selected locations in Great South Bay. New York.

Stony Brook, NY: State Univ. of New York. 199 p. Thesis.
Hazlett, B. A. 1962. Aspects of the biology of snapping shrimp

(Alpheus and Synalpheus). Crustaceana (Leiden) 4:82-83.
Hunt, J. H. 1981. The importance of adult-larval interactions in

determining abundance patterns of soft-sediment infauna.

Chapel Hill, NC: Univ. of North Carolina. 59 p. Thesis.
Landers, W. S. 1954. Notes on the predation of the hard clam, Venus

mercenaria, by the mud CTab.Neopanope taxana. Ecology 35:422.
Loosanoff, V. L. 1959. Condylostoma-zn enemy of bivalve larvae.

Science 129:147.
Mackenzie. C. L., Jr. 1977. Predation on hard clam (Mercenaria

mercenaria) populations. Trans. Am. Fish. Soc. 106:530-537.
Menzel. R. W. 1970. The species and distribution of quahog clams

Mercenaria. Proc. Natl. Shellfish. Assoc. 60:8 (abstract).
, E. W. Cake, M. L. Haines. R. E. Martin & L. A. Olsen.

1976. Clam mariculture in northwest Florida: field study on

predation. Proc. Natl. Shellfish. Assoc. 65:59-62.
Menzel, R. W. & H. W. Sims. 1964. Experimental farming of hard

clams, Mercenaria mercenaria. in Florida. Proc. Natl. Shellfish.

Assoc. 53:103-109.
Nolan, B. A. & M. Salmon. 1970. The behavior and ecology of

snapping shrimp (Crustacea: Alpheus heterochelis and Alpheus

normanni). Forma Functio 2:289-335.
Peterson. C. H. 1982. Clam predation by whelks (Busycon spp.):

Experimental tests of the importance of prey size, prey density

and seagrass cover. Mar. Biol. (Berl.) 66:159-170.
Schein, H. 1977. The role of snapping in Alpheus heterocliaelis Say,

1818, the big-clawed snapping shrimp. Crustaceana (Leiden)

33:183-188.
Sokal, R. R. & F. J. Rohlf. 1969. Biometry: The Principles and

Practice of Statistics in Biological Research. San Francisco, CA:

W. H. Freeman and Co.
Street, M. 1982. Trends in North Carolina's commercial fisheries,

1965-1981. NC Dep. Nat. Resour., Comm. Dev. Div. Mar. Fish.

17 p.
Vermeij. G. J. 1978. Biogeography and Adaptation: Patterns of

Marine Life. Cambridge, MA: Harvard Univ. Press.
Whetstone, J. M. & A. G. Eversole. 1978. Predation on hard clams.

Mercenaria mercenaria . by mud crabs, Panopeus herbstii. Proc.

Natl. Shellfish. Assoc. 68:42-48.

Journal of Shellfish Research, Vol. 3, No. 1, 11-17, 1983.

SEASONAL GONADAL DEVELOPMENT OF YOUNG LABORATORY-SPAWNED

SOUTHERN {MERCENARIA CAMPECHIENSIS) AND NORTHERN

(MERCENARIA MERCENARIA ) QUAHOGS AND THEIR

RECIPROCAL HYBRIDS IN NORTHWEST FLORIDA

RODNEY DALTON 1 AND WINSTON MENZEL

Department of Oceanography
Florida State University
Tallahassee, Florida 32306

ABSTRACT The seasonal gonadal development of laboratory-spawned southern and northern quahogs and their recipro-
cal hybrids was investigated. All young clams were males and one or more stages of gametogenic activity were seen each
month of the year. Winter spawning, which occurred in all pedigrees of quahogs, was considered abnormal and resulted
from the unusually warm winter of 1 9 74- 7 5. Gonadal development of the hybrid 9 Mercenaria campechiensis
X 6 Mercenaria mercenaria was similar to its southern parent; the reciprocal hybrid was similar to its northern parent.
This may indicate maternal influence. Little or no spawning by M. campechiensis during warmer months was unlike that of
the other three pedigrees. Temperature was the overall controlling factor in gonadal development and spawning, but
genetic differences existed between the two species.

KEY WORDS Genetics, gametogenesis, hybridization, hard clams, quahogs, Mercenaria spp.

INTRODUCTION

The seasonal gonadal development of the northern
quahog clam Mercenaria mercenaria (Linne) has been studied
from the New England area (Loosanoff 1937a.b), from
Delaware Bay (Keck et al. 1975), from North Carolina
(Porter 1964), and from South Carolina (Eversole et al.
1980). A closely related species, the southern quahog
Mercenaria campechiensis (Gmelin). hydridizes readily with
the northern quahog (Loosanoff 1954) and the hybrids are
fertile (Menzel and Menzel 1965, Menzel 1968), hut the
reproductive cycles of neither the southern nor the hybrids
have been investigated. The present study is of the seasonal
gonadal cycles of young, laboratory-spawned northern and
southern quahogs and their reciprocal hybrids cultured in
northwestern Florida. The results are compared with
published reports from other areas.

MATERIALS AND METHODS

Southern quahogs, previously collected in the vicinity of
Florida State University (FSU) Marine Laboratory, north-
western Florida, were spawned by Dr. Charles Epifanio at
the University of Delaware Center for Mariculture Research
on 2 April 1974. Wild northern quahogs from Delaware Bay
were also spawned. Besides making self-fertilizations of
each species, reciprocal hybrids between the species were
produced. The larvae were cultured to metamorphosis and
grown to a size of 1 to 2 mm before shipment to Florida in
late June 1 974. The clams were reared to a size of 4 to 8 mm
at the FSU Marine Laboratory. On 4 October 1974, they
were planted in 10-cm deep, sandfilled, screen-covered

Present address: National Marine Fisheries Service, 9450 Roger
Blvd., St. Petersburg, FL 33702

wooden boxes in Alligator Harbor, about 8 km from the
laboratory. At mean low water 4 to 5 cm of water covered
the clams.

Ten clams of each pedigree were sampled on the 5th
(± 1 day) of each month from 6 November 1974 through
5 November 1975, and additional samples were taken on
the 20th (± 1 day) in December 1974, and in September
and October 1975. The total sample included 660 clams
from which 6,000+ follicles were microscopically examined.
After February 1975, the stock of the hybrid 9 Mercenaria
mercenaria X 6 Mercenaria campechiensis was depleted,
primarily from crab predation. Additional clams of the
same pedigree, planted as surplus in the same area, were
sampled from May 1975 until the stock became exhausted
by August 1975.

Shucked clams were preserved in Bouin's fixative, trans-
ferred to alcohol, imbedded in Paraplast®, sectioned at
8 nm, mounted on slides, and stained with Erlich's hemo-
toxylin and erosin following standard histological proce-
dures. Previous examinations showed that transverse mid-
longitudinal sections gave a good representation of the
gonad condition. All follicles in the most representative of
8 to 10 sections of each clam were used to determine
gonadal condition.

Determination of gonadal condition followed that of
Ropes (1968) as modified by Haines (1976). As noted by
Loosanoff (1937a). different follicles within the same clam
and different clams within the same population were often
in several stages of gonadal development. The gonadal stages
are not illustrated because they have been reported pre-
viously by Loosanoff (1937a), Porter (1964), Keck et al.
(1975), and Eversole et al. (1980). Brief descriptions of
each stage follow.

Dalton and Menzel

Indifferent or Spent

The lumen of the indifferent or spent follicles are usually
conspicuously empty, although a few residual spermatozoa
may be present (in spent follicles) and a few scattered
spermatogonia occur around the membranes of the other-
wise bare follicles.

Early to Late Active

Follicles in the early active stage are undergoing primary
and secondary spermatogenesis, with a nearly continuous
layer of cells forming around the follicle membrane. Later,
the lumen fills with basophilic spermatids and a few sperma-
togonia occur near the periphery. Early and late active
stages were recorded separately but are presented as active
stage only.

Ripe

The ripe phase is easily distinguished by a dense mass of
spermatozoa, filling the follicles. Other types of gameto-
genic cells may be present, but are not abundant.

Partially Spent

Partially spent follicles contain spermatozoa within the
lumen of the follicle but these are substantially less abun-
dant than in the ripe stage.

Percentages of each gonadal stage for each pedigree at
each sampling were graphed and the mean percentages of
each stage of each pedigree were calculated and graphed
to emphasize the similarities and differences between the
four pedigrees. The first samples (November 1974) were
not included in the mean calculations because no clams
were mature enough to spawn and the results would be
biased. Additionally, because of the smaller amount of data
for the hybrid 9 Mercenaria mercenaria X d Mercenaria
campechiensis, comparative data were recalculated using
only samples collected in November 1974-February 1975,
and May-August 1975.

Water temperatures were taken at time of sampling at
depths of 20 to 30 cm. These infrequent observations were
supplemented with minimum and maximum air tempera-
tures (mean of 6-day intervals) from local climatological
data recorded at Apalachicola, FL (NOAA 1974a, 1975a).
Although Apalachicola is about 50 km from Alligator
Harbor, that coastal location has the same latitude and is
considered representative for this study.

In April 1976, when the clams were two years old, the
remaining 19 southern, 4 northern and 11 hybrids
(9 Mercenaria campechiensis X 6 Mercenaria mercenaria)
were recovered and their sex was determined by the smear
technique.

RESULTS

Both of the species and the hybrids were predominantly
male. Two clams (0.3%) showed evidence of oogenesis.
The follicles were in the early active stage, but no clams

were observed with ripe female follicles. Occasionally, a few
early stage female gamete cells occurred in otherwise male
follicles, indicating a possibility for hermaphroditism. Game-
togenesis had commenced by the first examination in Novem-
ber 1974, when the quahogs were seven months old, but
only 2 to 4 follicles were seen per histological section. Later,
the number of follicles increased to 15 to 20 per section.
Gametogenesis in one or more stages were seen throughout
the entire period in all the samples and pedigrees. Differences
in the seasonal occurrence and relative overall abundance of
each stage occurred in each pedigree. A discussion of the
seasonal occurrence of each gonadal stage and probable
times of spawnings are given for each pedigree.

Southern Quahog, Mercenaria campechiensis

Indifferent or spent follicles were present in all the
samples of the southern quahog (Figure 1 ) and were in the
largest mean percentage, 54% (Figure 2A). Active stages
were also seen in all the samples except that taken 5 April,
but occurred in low percentages in December, May, and
June, with values of 10, 5, and 6%, respectively (Figure 1).
The mean percentage for the entire period was 23%
(Figure 2A). The percentages of ripe stage follicles were
highest in both samples taken in December (47% and 40%)
and in January (43%). This stage decreased in February
(10%), March (14%), and April (6%), and none or very low
percentages occurred through the 20 September sample
(6%). Ripe follicles were found in the remaining samples
(9-14%) (Figure 1). The mean for the entire period was
13% (Figure 2 A). Partially spent stages were first seen in
the sample taken 20 December (9%) and continued in
relatively high percentages through the 5 April period
(10—32%). This stage decreased by the May sample (6%)
and was low until the following fall, increasing to 17% on
5 October (Figure 1 ). The mean was 10% (Figure 2A).

Spawning, as indicated by comparison of ripe and
partially spent stages, commenced after the 5 December
sample and continued until 5 April, with a probable peak in
March. Little or no spawning occurred during the summer
months, but spawning commenced again after 5 September.

Northern Quahog, Mercenaria mercenaiia

Indifferent or spent follicles were present in all samples
of Mercenaria mercenaria (Figure 1) but in considerably
less abundance (X = 28%) than for the southern species
(Figure 2A). Active stage follicles were also present in all
samples (X = 58%) and in greater abundance than the
southern species (Figure 2A). Ripe follicles occurred in all
sampling periods, except the first on 6 November and
those on 5 May and 20 September (Figure 1 ) (X = 10%)
(Figure 2A). Partially spent follicles were seen in the samples
taken 5 and 20 December, but not again until 5 March,
when the highest percentage occurred (13%). This stage
occurred on all the other sampling dates except that taken
on 5 May (Figure 1 ). The mean was 4% (Figure 2A).

Gonadal Di vi lopmtNt oi young Quahogs

Mercenaria campechiensis

Mercenaria mercenaria

DJFMAMJJAS ON

N D JFMAMJJA S N

^Mercenaria campechiensis

X
^Mercenaria mercenaria

N D JFMAMJJA S

CH Indifferent/Spent
ES3 Active

^Mercenaria mercenaria

X
dMercernaria campechiensis

ON N D J F

Months
Ripe
] Partially Spent

nAAA/

M J J A

Figure 1. Reproductive cycles of southern and northern quahogs and their hybrids (660 total) shown as the percentage of
follicles (males only) in each gonadal stage (period from 6 November 1974 through 5 November 1975).

Dalton and Menzel

ru-

60-

50-

40-

30-

20-

10-

Mm 9Mc x cfMm

Indifferent/Spent ^SB Ripe

C*3*l Active r^x^l Partially Spent

9Mc x cfMm 9Mm x cfMc

Figure 2. Mean percentages of follicle stages in southern (Mc) and northern (Mn) quahogs and their hybrids. (A) December 1974-November
1975: southern, northern and 9 southern X d northern. (B) December 1974-Februaiy 1975 and May-August 1975: southern, northern
and reciprocal hybrids.

The data for ripe and partially spent follicles indicate
that spawning started by 5 December, but ceased from
20 December until after the 5 February sample. A peak of
spawning occurred between 5 February and 5 May, with a
probable high in March. Spawning resumed after 5 May and
continued throughout the balance of the sampling period;
a probable secondary peak occurred in September.

Hybrid, 9 Mercenaria campechiensis X d Mercenaria mercenaria

The sequences of follicle development stages in the hybrid
9 Mercenaria campechienses X 6 Mercenaria mercenaria are
similar to the southern quahog parent. Indifferent or spent
stages were found in all the samples (Figure 1 ) and. as in
M. campechienses, had the highest mean (58%) (Figure 2 A).
Active follicle stages were also present in all the samples,
ranging from a high of 54% on 20 December to lows of
17% in April, June, and July (Figure 1 ); the mean for the
entire period was 27% (Figure 2A). This hybrid was the
only pedigree that had ripe follicles (21%) on the first
sampling (6 November 1974). The highest percentages of
the ripe stage occurred on 5 January (34%) and on 5 March
(269! ). Ripe follicles were not seen in the 5 April samples
but were observed in varying percentages for the balance of
the sampling dates (Figure 1 ). The mean of the ripe follicles
was 10%' (Figure 2 A). Partially spent stages were first seen
20 December and continued through the 5 March sample;
none occurred on 5 April. This stage occurred in low
percentages for the balance of the period, except for none
on 20 December (Figure 1 ). The mean was 5% (Figure 2A).

The data indicate that spawning commenced after
5 December and continued through March. The absence of
both ripe and partially spent stages in the 5 April sample

indicates a peak of spawning in March. Spawning resumed
after 5 April and continued throughout the balance of the
examinations, with probable peaks in May-July and again
in September.

Hybrid, 9 Mercenaria mercenaria X d Mercenaria campechiensis

Unfortunately data for the hybrid 9 Mercenaria
mercenaria X d Mercenaria campechiensis are incomplete,
but those obtained show the sequences of follicle develop-
ment to be similar to the northern quahog. Indifferent or
spent stages were present in all the samples and ranged from
a high of 40%' on 5 June to a low of 5% on 5 December
(Figure 1) (X= 23%, Figure 2B). Ripe follicles (4%) first
seen on 5 December, increased to a high of 26%' on 5 Janu-
ary, and were found on all the other dates for which data
are available; another high (28%) occurred on 5 July
(Figure 1). The mean for the entire period was 15%
(Figure 2B). Partially spent follicles were first observed on
20 December and were seen in all the other samples, except
that on 5 May (Figure 1 );X= 7% (Figure 2B).

Spawning commenced after 5 December and continued
to at least 5 February. The absence of partially spent
follicles on 5 May indicates that a peak of spawning occurred
prior to this date. Spawning continued after 5 May to at
least 5 August, the last date sampled.

Sex could be determined for only 15 of the 34 two-year-
old clams collected in April 1976. Of these clams. 13 were
males and 2 were females (2 of 4 northern sampled).

DISCUSSION

This is the first study of the seasonal gonadal develop-
ment of the southern quahog Mercenaria campechiensis and

Gonadal Development of Young quahogs

its hybrids with the northern species Mercenaria mercenaria,
with a comparison of laboratory-spawned clams of known
age grown in the semitropical area of northern Florida. This
study is not as thorough as those from more northern
latitudes because observations were made for only one year
and of male clams only. The spawnings that occurred in
the winter period were undoubtedly atypical and are
discussed in more detail below.

Loosanoff (1937a) found that quahogs have a protandric
development; almost all clams (98%) developed first as
males, but eventually achieved an equal sex ratio as older
clams. Eversole et al. (1980) also found a preponderance of
males to females (9.5:1) in young quahogs and a 1:1 sex
ratio in older animals. Our study confirms the protandric
development in northern quahogs and documents the same
type of development in the southern species and its hybrids.
The samples of 2-year-old clams revealed that sex reversal
to female was occurring, even though the sampling was
very small. Large clams of both species and hybrids that
were used in our spawning experiments over the past 20
years usually had a 1 : 1 sex ratio.

Only 2 to 4 follicles were present in the first sample
(6 November 1974) and were localized near the stomach
ventral of the pericardial sinus. This was the same location
reported by Loosanoff (1937a), but he found 6 to 8
follicles in clams of approximately the same size and
probably of lesser age. The slighter gonadal development of
quahogs grown in Florida was surprising, especially as growth
rates have been reported to be greater than in more northern
areas (Menzel 1961, 1962. 1977). One possible explanation
is that the animals were laboratory reared and cultured in
the natural habitat for only one month when first examined.
Growth has always been less under our laboratory condi-
tions than when planted in the open waters. Enough food
may have been available for shell growth but not enough
for gonadal development. Sastry (1966) stated that the bay
scallop Argopecten irradians (Lamarck) "requires large
amounts of food for gonad growth." Loosanoff and Davis
(1950) found that Crassostrea virginica (Gmelin) did not
mature sexually with poor glycogen reserve.

Figures 1 and 2, especially 2, show a usually low per-
centage of the partially spent stage in all the pedigrees.
This probably indicates that once spawning is initiated in
ripe clams, it is completed in a short period of time. If
partially spent follicles occur for only a brief period,
errors may have been made in deducing times of spawning,
which were based on comparisons of ripe and partially
spent clams at each examination (1 month inmost instances).

Spawning throughout the year in marine invertebrates
occurs most commonly in areas where there is little seasonal
change, such as the tropics, polar regions, and deep sea
(Goodbody 1965, Sanders and Hessler 1969). Northwestern
Florida is subtropical, but warmer than normal tempera-
tures occurred during the winter of 1975-75. Northern
Florida experiences periods of air temperatures below

freezing and water temperatures below 10°C;water tempera-
tures in January-February 1958—61 were as low as 6 to 9°C
(Menzel 1961). The lowest water temperature during the
winter of 1974-75 was 1 1.5°C in early December and air
temperatures at Apalachicola never dropped below freezing
(Figure 3). Extended periods occurred during the winter
of 1974—75 when air temperatures were above 20°C in
December-February (Figure 3). Those periods coincided
with minus spring tides of -5 to -40 cm during the hours
of 0730—1700 (National Oceanic and Atmospheric Admin-
istration, 1974b, 1975b). We have repeatedly observed in
our laboratory that when alternating thermal stimulation is
used to induce spawning, quahogs initiate spawning on the
decreasing temperatures. Also, males usually spawn before
females. The male quahogs in the boxes may, therefore,
have been warmed to the critical spawning temperatures
during the minus tides on warm days and stimulated to
spawn when covered by the cooler incoming water at
flood tide.

All quahog pedigrees had ripe follicles during winter
months. This is consistent with other observations. Chestnut
(195 1) found that Mercenaria mercenaria often reach sexual
maturity by mid-winter in North Carolina. Our thermal-
induced laboratory spawning of both sexes has been most
successful during the winter months. Winter spawnings are
unusual in northern Florida. All wild quahogs have been
found subtidally; a few may be uncovered by low tides of
> —30 cm. Even if winter spawning does occur, it is unlikely
that the gametes/larvae would survive in the relatively cold
water. A larger percentage of the follicles may have been in
the ripe condition during the winter months if normal
temperatures had prevented spawning.

Reproductive cycles in marine invertebrates vary with the
latitude and modifications have been associated with differ-
ences in temperature regimes (Orton 1920. Nelson 1928,
Thorson 1950, Loosanoff and Nomejko 1951, Sastry and
Blake 1971). The northern quahog ranges from Canada
southward on the Atlantic coast and throughout the
northern Gulf of Mexico (Abbot 1974) and thus experiences
a wide range of temperatures. The spawning periods of the
northern quahog have been documented for the areas
ranging from Long Island Sound to South Carolina and now
for northern Florida. The spawning periods in Florida,
disregarding the winter spawning, showed bimodal spawning
peaks in the spring and fall similar to that observed in the
Carolinas (Porter 1964, Eversole et al. 1980); however,
spawning began about a month (March) earlier and extended
about a month (October) later than in the Carolinas. These
northern clams were the progeny of clams native to Delaware
Bay, where there is a single peak of spawning (Keck et al.
1975), similar to Long Island Sound (Loosanoff 1937b).
Peak spawnings by southern and northern quahogs and the
reciprocal hybrids were essentially the same.

We noted that percentages of indifferent/spent and active
stages of gonadal activity of the southern species and the

Dalton and Menzel

35
30
25

♦"

(MAX)

(MIN)

1974

1975

Figure 3. Water temperatures (heavy line) at Alligator Harbor and maximum and minimum air temperatures (mean of 6-day intervals) at
Apalachicola, Florida.

hybrid 9 Mercenaria campechiensis X 6 Mercenaria
mercenaria were very similar; whereas, the northern and the
other hybrid were similar. Menzel (1962) has reported that
hybrid quahogs in Florida grew faster than their northern
parents and were more like the faster growing southern
parent. The hybrid 9 M. campechiensis X 6 M. mercenaria
had a slightly better growth rate than the reciprocal hybrid
indicating the possibility of maternal influence.

It would be interesting to determine the seasonal gonadal
development of females of both species and hybrids in
Florida. Previous observations in our laboratory have shown
that it is virtually impossible to induce summer spawning of
females of any pedigree after about March-April when the
ambient water temperatures exceed 22 to 24°C. Active
sperm appear in suspensions but few ripe ova occur in clams
during the warmer months. Successful female spawnings
have been induced during periods from October-March with

no temperature conditioning. The seasonal gonadal
development, therefore, may be different for female
quahogs than reported here for young males.

Also, it would be interesting to determine if quahogs of
both species follow the pattern of gametogensis of the
endemic population when transplanted to a colder latitude.
Such observations might be difficult because the southern
quahog and the hybrids lack a tolerance to low tempera-
tures (Chestnut et al. 1956, Haven and Andrews 1956,
Menzel 1977). Whether the northern quahog, native to
warmer areas, would survive in cold winter regions is not
known. Belding (1912) reported 70 years ago that tempera-
ture is the controlling factor in quahog spawning. Based
on the data of all the investigations, we believe that both
species and the hybrids will have generally similar gamete
development and spawning, regardless of their origin,
within a specific area.

Gonadal Development of Young Quahogs

REFERENCES CITED

Abbott, R. T. 1974. American Seashells. New York, NY: Van

Reinhold Company. 2nd edition. 663 p.
Belding, D. L. 1912. A report upon the quahog and oyster fisheries

of Massachusetts, including the life history, growth and cultiva-
tion of the quahog (Venus mercenaria), and observations on the

set of oyster spat in Well Fleet Bay, Boston. Boston, MA: Wright

and Potter Print Co. 134 p. (Reissued: 1964. Mass. Dep. Nat.

Resour. Div. Mar. Fish., Contrib. 12:134 p.)
Chestnut, A. F. 1951. The oyster and other mollusks in North

Carolina. Taylor, H.F., ed., Survey of Marine Fisheries of North

Carolina. Chapel Hill, NC: Univ. N.C. Press; 141-190.
, W. E. Fahy & H. J. Porter. 1956. Growth of young Venus

mercenaria. Venus campechiensis, and their hybrids. Proc. Natl.

Shellfish. Assoc. 47:50-56.
Eversole, A. G., W. K. Michener & P. J. Eldridge. 1980. Reproductive

cycle of Mercenaria mercenaria in a South Carolina estuary. Proc.

Natl. Shellfish. Assoc. 70:22-30.
Goodbody, I. 1965. Continuous breeding in populations of tropical

crustaceans, Mysidium columbiae (Zimmer) and Emerita portori-

censis (Schmidt). Ecology 46:195-197.
Haines, M. L. 1976. The reproductive cycle of the sunray venus

clam, Macrocallista nimbosa (Lightfoot, 1786). Proc. Natl.

Shellfish. Assoc. 66:6-12.
Haven, D. & J. D. Andrews. 1956. Survival and growth of Venus

mercenaria, Venus campechiensis, and their hybrids in suspended

trays and on natural bottoms. Proc. Natl. Shellfish. Assoc.

47:43-49.
Keck, R. T., D. Maurer & C. H. Lind. 1975. A comparative study of

the hard clam gonad developmental cycle. Biol. Bull. (Woods

Hole) 148:243-258.
Loosanoff, V. L. 1937a. Development of the primary gonad and

sexual phases in Venus mercenaria Linnaeus. Biol. Bull. (Woods

Hole) 72:389-405.
. 1937b. Seasonal gonadal changes of adult clams, Venus

mercenaria (L.). Biol. Bull. (Woods Hole) 72:406-416.
. 1954. New advances in the study of bivalve larvae. A m. Sci.

43:607-624.
& H. C. Davis. 1950. Conditioning Venus mercenaria for

spawning in winter and breeding its larvae in the laboratory. Biol.
Bull. (Woods Hole) 98:60-65.
Loosanoff, V. L. &C. A. Nomejko. 1951. Existence of physiologically
different races of oyster, Crassostrea virginica. Biol. Bull. (Woods
Hole) 101:151-156.

Menzel, R. W. 1961. Seasonal growth of the northern quahog,
Mercenaria mercenaria and the southern quahog,M campechiensis,
in Alligator Harbor, Florida. Proc. Natl. Shellfish. Assoc. 52:
37-46.

. 1962. Seasonal growth of the northern and southern

quahogs, Mercenaria mercenaria and M. campechiensis, and their
hybrids in Florida. Proc. Natl. Shellfish. Assoc. 53:111-119.

. 1968. Cytotaxonomy of species of clams (Mercenaria)

and oysters (Crassostrea). Symp. Mollusca, Mar. Biol. Assoc.
India. Part 1:75-84.

. 1977. Selection and hybridization in quahog clams

(Mercenaria spp.). Proc. World Maricult. Soc. 8:507-521.
& M. Y. Menzel. 1965. Chromosomes of two species of

quahogs and their hybrids. Biol. Bull. (Woods Hole) 129:

181-188.
Nelson, T. C. 1928. On the critical temperatures for the spawning

and for ciliary activity in bivalve molluscs. Science 67:220-221.
National Oceanic and Atmospheric Administration. 1974a. Climato-

logical Data, Florida. U.S. Dept. Commerce. 78.
. 1974b. Tide Tables, East Coast of North and South

America. U.S. Dept. Commerce.
. 1975a. Climatological Data, Florida. U.S. Dept. Com-

merce. 79.

. 1975b. Tide Tables, East Coast of North and South

America. U.S. Dept. Commerce.
Orton, J. H. 1920. Sea temperature, breeding and distribution in

marine animals. J. Mar. Biol. Assoc. U.K. 12:339-366.
Porter, H. J. 1964. Seasonal gonadal changes of adult clams,

Mercenaria mercenaria (L.) inNorth Carolina. Proc. Natl. Shellfish.

Assoc. 55:35-5 2.
Ropes, J. W. 1968. Reproductive cycle of the surf clam, Spisula

solidissima. in offshore New Jersey. Biol. Bull. (Woods Hole)

135:349-365.
Sanders, H. L. & R. R. Hessler. 1969. Ecology of the deepsea

benthos. Science 163:1419-1424.
Sastry, A. N. 1966. Temperature effects in reproduction of the bay

scallop, Aequipecten irradians Lamarck. Biol. Bull. (Woods

Hole) 130:118-134.
& N. J. Blake. 1971. Regulation of gonad development

in the bay scallop, Aequipecten irradians Lamarck. Biol. Bull.

(Woods Hole) 140:274-283.
Thorson, G. 1950. Reproductive and larval ecology of marine

bottom invertebrates. Biol. Rev. Camb. Philos. Soc. 25:1-45.

Journal of Shellfish Research, Vol. 3, No. 1, 19-27, 1983.

EXPERIMENTAL PLANTINGS OF JUVENILES OF THE HARD CLAM

MERCENARIA MERCENARIA (LINNE) IN THE WATERS OF

LONG ISLAND, NEW YORK 1

PAUL J. FLAGG AND ROBERT E. MALOUF

Marine Sciences Research Center
Stare University of New York
Stony Brook, New York 11794

ABSTRACT Planting of hatchery-reared seed of the hard clam Mercenaria mercenaria is a significant management tool
in town-managed shellfisheries of New York. In the present study, seed planting techniques developed elsewhere were
tested in New York waters. The objectives were to determine how seed survival was influenced by ( 1) seed size at the time
of planting; (2) the presence, absence, and type of gravel aggregate; (3) the season planted; and (4) site selection. Site
characteristics, particularly the types and abundance of predators present, were found to influence the results so strongly
that general recommendations cannot be made. Mud crabs (Neopanope sayi [Smith] ) and whelks (Busy con carica [Gmelin]
and B. canaliculatum [Linne]) were the most damaging predators at the sites tested. Gravel aggregate did not provide
adequate protection for planted clams, and the use of large (25-mm) gravel appeared to have a negative impact on seed
survival. Survival exceeded 10% only among clams that were at least 20 mm in length at planting; however, mortalities
as high as 100% resulted from plantings of such seed (23 mm) at sites having significant populations of whelks.

KEY WORDS: Hard clams, Mercenaria mercenaria, seed planting, predation

INTRODUCTION

The hard clam (or northern quahog) Mercenaria
mercenaria (Linne) is the object of New York's most
important shellfishery, accounting in recent years for
about 50% of the total value of fishery products landed in
the state (McHugh and Ginter 1978). Long Island's Great
South Bay is the single most important producer of hard
clams in the world. This 24,282-ha (60,000-acre) bay has
historically produced about 90% of the New York harvest
and 45% of the total United States harvest of hard clams.
Since 1977, New York landings of hard clams have declined
dramatically. For example, the 1976 reported Great South
Bay landings were 24,684 m 3 (700,465 bu), but by 1981,
the landings had dropped to 10,758 m 3 (305,287 bu)
(National Marine Fisheries Service, Patchogue, NY, unpub-
lished fishery statistics, 1982).

Although stock assessment data are incomplete, declining
harvests are perceived by many local fishery managers to
represent a real drop in standing stocks (J. Kassner, Town
of Brookhaven, NY, and Pieter Van Volkenburgh, NY Dept.
Environm. Conserv., Stony Brook. NY, pers. comm.). Local
management agencies, primarily the townships, have
responded to declining landings by instituting programs
intended to supplement natural hard clam reproduction.
Among the most popular programs are those that involve
the planting of seed clams. Nine Long Island townships,
including all three of the townships that border Great South
Bay. have carried out some type of seed clam planting
program. Their efforts have ranged from trial plantings of a

Contribution No. 378 of the Marine Sciences Research Center,
State University of New York (SUNY) at Stony Brook.

few thousand seed to annual plantings in excess of 1 million
seed. Seed are purchased from a commercial hatchery,
held in some type of nursery system, and eventually broad-
cast onto the bay bottom without any protection. Nursery
systems used include shore-based raceways and ponds,
rafts, and gravel beds. The size of the seed at the time of
release to the public fishery generally ranges from about
8 to 25 mm in shell length.

There are no published studies of seed clam plantings in
New York waters. In fact, some doubt has been expressed
that the seed planting programs can possibly be of sufficient
scale to significantly impact the fishery (McHugh 1981).
The early work of Haven and Andrews (1957) showed that
seed clams require some type of protection to ensure survival.
Similarly, Menzel and Sims ( 1964) reported that seed clams
planted in Florida required protection or had to be at least
12 mm in shell length to avoid very heavy predation losses.
Castagna (1970) demonstrated that gravel aggregate helped
prevent the loss of seed clams. Castagna and Kraeuter ( 1977)
and Kraeuter and Castagna (1977) recommended the use of
aggregate as part of a culture system that included baffles
and fences. Their work and the work of Menzel et al.
(1976) suggested that the use of stone aggregate alone
affords planted seed clams some protection from predators.
The use of stone aggregate would be particularly attractive
for the extensive nursery plots that are required for large
public fisheries because of its relative simplicity and low
cost; it has been used on a limited basis for that purpose
(Jeffrey Kassner, Town of Brookhaven, NY. pers. comm.).

Eldridge et al. (1979) made the following recommenda-
tions based on several years of seed clam planting in South
Carolina: (1) select a physically suitable habitat, one that

Flagg and Malouf

is free, for example, from extreme wave action; (2) cover
the planting area with shell or stone aggregate; (3) plant
seed clams in the fall when temperatures are 15 to 18°C;
(4) plant seed of 12 to 15 mm shell length at a density of
300 m 2 ; and (5) harvest in the early summer of the second
year. The authors pointed out that uncontrolled variables
contribute to the uncertainty of such a planting as a private
venture; however, they reported approximately 77% annual
survival of 16- to 17-mm seed and 95% annual survival of
21- to 22-mm seed planted in this manner. Later work by
Whetstone and Eversole (1981) also reinforced the case
for fall plantings by demonstrating in laboratory studies
that the activity of an important hard clam predator, the
common mud crab Panopeus herbstii H. Milne-Edwards,
was significantly reduced at temperatures below 17°C.

The present study was part of an effort to test and
refine a number of seed-clam planting techniques that
have been developed elsewhere. The intention was to
evaluate recommended planting procedures for possible
application to a large public clam fishery. Specifically, the
objectives were to determine in New York waters how the
survival of three sizes of planted seed clams was affected:
(1) by the size and shape of aggregate and sand substrate
(Experiment I); (2) by the time (season) they were planted
and recovered (Experiment II); and, (3) by site specific
environmental differences within the same general location
(Experiment III).

materials and methods

Experiment I was sited in a shallow cove, separated by a
sand spit from Eastern Shinnecock Bay, Long Island, NY
(designated as Site I, Figure 1 ). Mean low water depth at
the site was approximately 0.5 m, and the tidal range
averaged about 1.0 m. Sediments within the cove graded
from coarse sand near the sand bar to soft mud near the
northern edge of the cove. Eeel grass (Zostera marina
Linnaeus) was present, but was relatively sparse through
most of the planting area. A natural population of adults of
Mercenaria mercenaria existed in the cove prior to our
planting at a mean density of about 7 clams m" 2 .

The seven substrates tested in this experiment consisted
of sand and two shapes of gravel obtained in three sizes.
The two shapes were (1) mechanically produced, crushed
gravel having irregular shapes and jagged edges, and (2)
more rounded, unbroken glacial gravel. Both gravel types
were obtained in three nominal sizes: 6 to 10, 10 to 19, and
19 to 32 mm. The gravel was washed through wire screens
to obtain the approximate size ranges given above. All
gravel was obtained from Long Island glacial till and was
washed thoroughly with fresh water during processing.

Forty-two plastic, food-handling trays (Nestier® "Chill-
tray 180") measuring 56.5 X 46.4 X 17.8 cm) were lined
with 2-mm mesh plastic window screen. The trays were
filled to a depth of approximately 8 cm with 20-mm gravel.
They were then transported to the site, arranged in a

6X7 array, and hydraulically sunk (jetted) into the bottom
so that approximately 3 cm of the tray edges protruded
above the substrate. A 4-cm layer of one of the seven types
of substrate was then added to the surface of each tray in a
randomly generated pattern.

Three sizes of seed clams used in the experimental
plantings were obtained from Aquaculture Research Corp.,
Dennis, MA. At the time of planting (23 July 1980), the
mean shell lengths and standard errors (n = 50) for clams
of the size groups were 3.9 ± 0.06, 6.8 ± 0.08, and 28.7 ±
0.23 mm. Planting densities used were 1,241, 477, and
191 m" 2 for the small, medium, and large seed, respectively.
Thus, a tray randomly received 325 small, 125 medium, or
50 large seed. The experimental design included two
replicate plantings for each treatment. Because there was
no differentiation of substrate shape for plantings in sand,
for each clam size there were four replicate plantings in
sand. Also, because they were in short supply, the largest
seed clams were only planted in the three sizes of round
gravel and in sand.

The planting area was examined weekly to identify and
count potential clam predators. The experiment was
terminated on 20-22 October, when water temperatures
in the area dropped below 10°C. The trays were lifted on
board a small boat, and all remaining clams were removed
and counted and their shell lengths were measured to the
nearest millimeter. Empty shells and shell fragments were
examined for evidence of predation, and any predators
recovered with the trays were identified and counted.

Growth and survival (recovery) data were statistically
analyzed by analysis of variance ( ANOVA) following Sokal
and Rohlf (1969). Shell length measurements were used to
calculate growth in millimeters.

Experiment II was initiated in the fall of 1980 at two
locations (designated Sites IIA and IIB, Figure 1) in Eastern
Long Island. Site IIA was located in Shinnecock Bay
approximately 30 m east of the previously described site
of Experiment I. Site IIA had a mean low water depth of
approximately 0.35 m, and bare sandy sediments. Site IIB
was located in Napeague Harbor, Long Island. Mean low
water depth at the site was 1.0 m, and the tidal range was
0.9 m. Sediment at Site IIB consisted of a 3-cm-deep layer
of sand over gravel and stones. The area was devoid of eel
grass and macroalgal detritus. A sparse (< 1 irf 2 ) natural
population of very large hard clams existed at Site IIB prior
to our planting.

Experiment II consisted of two replicate plantings of
each of three clam sizes in two substrates types (sand and
1 cm crushed gravel) at two sites and at two planting times.
The two planting times and ambient water temperatures at
the two sites were: 30 September 1 980 ( 1 9°C) and 25 Novem-
ber (8°C) for Site IIA, and 30 September 1980 (17°C) and
22 November 1980 (8°C) for Site IIB. Seed clams were
again purchased from Aquaculture Research Corp. Mean
shell lengths and standard errors (n = 50) for the three size

Experimental Plantings of mercenaria mercenaria

CT.

10 n mi
-I
5km

.«&cP

ATLANTIC OCEAN

Shmnecock
Bay

meters

I 1 1

1000

ATLANTIC OCEAN

Nope ague
Bay

meters

ATLANTIC OCEAN

Figure I. Location of six sites used for experimental plantings of seed clams on the south shore of Long Island, New York.

FLAGG AND MAI.OUF

classes in the September planting were 2.8 ± 0.17, 7.1 ±
0.10, and 22.7 ± 0.15 mm. Rapidly declining ambient water
temperatures necessitated the planting of the November
shipment immediately upon receipt. Therefore, although
hatchery sorting through sieves was identical for the two
shipments, shell measurements for the November shipment
were not recorded. Tray handling, seed-planting procedures,
and planting densities were as in Experiment I.

The planting sites were inspected regularly for predator
distribution and abundance. Final sampling of the trays
was conducted 9 months after the planting date (15— 22 June
and 23—27 August 1981 for the September and November
plantings, respectively). Sampling procedures and data
analysis were as in Experiment I except that no growth
analyses were included in Experiment II.

Experiment III consisted of plantings on prepared natural
bottom without trays. Plantings were carried out at three
sites (designated as Sites IIIA, IIIB, and IIIC, Figure 1 ) in
one general location, Napeague Harbor, Long Island.
Three sizes of seed clams (nominally, 3, 6, and 23 mm in
length) were planted at each site, with and without gravel,
during the summer of 1 98 1 .

Site IIIA was located approximately 40 m east of Site
IIB, described above. The site had a mean low water depth
of 1.2 m and a tidal range of 0.9 m, and contained poorly
sorted sand and gravel sediments.

Site IIIB, in northeastern Napeague Harbor, had a mean
low water depth of 0.4 m and a tidal range of 0.9 m.
Sediments at the site consisted of coarse sand sparsely
interspersed with rocks. The site was on the edge of an
approximately 1 ha bare area in an eel grass flat. A dense
(20 to 50 rrf 2 ) population of small adult hard clams existed
at the site prior to our planting.

Site IIIC was located on a large bare sand/mud flat in the
southwestern part of the harbor. Mean low water depth was
0.4 m and tidal range was 0.9 m. Hard clams, predominately
adults plus a few subadults, were moderately abundant
(5 to 10 irf 2 ) prior to our planting.

Seed clams were purchased from the same commercial
source in the same three nominal sizes as used in the pre-
viously described experiments (2 to 4, 6 to 8, and 22 to
28 mm length). Each of the three sites consisted of six
2- X 2-m subsites delineated by 30- wide X 15-cm-deep
borders of 3-cm gravel. Each of the three seed clam sizes
were randomly assigned to two subsites. One of the two
subsites contained existing substrate, while the other
contained a 2.5-cm-deep layer of 1 .0 cm gravel. On 20 May
1981, clams were planted at all sites at densities of 1,250,
675, and 260 m~ 2 for small, medium, and large clams,
respectively.

Surveys of predator abundance were conducted prior to
planting (17-20 May 1981) and were repeated on 26—28
July and 13—14 September 1981. Sampling areas adjacent
to each site (30 m 2 in May and July and 15 m 2 in Septem-
ber) were raked with a clam rake lined with 1.3-cm Vexar®,

and predators were collected, counted, and measured. Esti-
mates of the abundance of the more mobile crabs (primarily
Ovalipes ocellatus [Herbst] ) were subject to error because
of the animals' mobility and are, therefore, not quantitative.
Sampling to determine seed clam survival was conducted
approximately two months after planting (26 July) and
again at termination (14 September). For purposes of
sampling, each subsite was divided into four 1 m 2 quadrats
and each quadrat into nine equal parts (0.1 1 m 2 each). Two
of the 0.1 1 m 2 areas were randomly selected from each of
two randomly selected quadrats. A 0.10 m 2 sampling
square was placed on a selected area, and substrate was
removed to a depth of 15 cm. After being separated from
the substrate, surviving clams were counted and returned
to the sample area. Analysis of survival data was as described
above.

RESULTS

Experiment I

The most abundant clam predators observed in and
around the trays following planting were Say's mud crabs
(Neopanope sayi), calico crabs {Ovalipes ocellatus),
channeled whelks (Busycon canaliculatwn), and oyster
drills (Urosalpinx cinerea [Say] and Hupleura caudata [Say] .
Other potential predators which were less frequently
observed included blue crabs (Callinectes sapidus Rathbun),
common mud crabs (Panopeus herbstii), and both winter and
summer flounders (Pseudopleuronectes americans [Wal-
baum] and Paralichthys dentatus [Linnaeus]), respectively.

The abundance of the mud crab N. sayi was positively
related to increased gravel size (Table 1). Those trays filled
with 19- to 32-mm gravel contained numerous 0-year-class
crabs. Up to 10 oyster drills (U. cinerea and E. caudata)
per tray occurred during the summer, but no drills were
found in the trays during the autumn sampling. Similarly,
channeled whelks (B. canaliculatum) were visible at the
substrate surface, and were most abundant during the
first month (August) following planting. Few were observed
later in the summer, and only two were recovered from the
trays during sampling.

Survival (recovery ) of planted seed clams was significantly
influenced by their size at the time of planting (0.01 >?>
0.001). Mean survival rates for small, medium, and large
clams were 4.0, 43.1, and 82.5%, respectively. The size of
the gravel used also significantly affected clam survival
(0.01 > P > 0.001). Further, the relationship between grain
size, independent of shape, and clam survival appeared to
be related to clam size (the interaction was significant;
P < 0.01 ). The smallest seed clams planted (4 mm) did not
survive well under any conditions. On the other hand, the
survival of the 29-mm seed was high and was independent
of grain size. The influence of grain size on clam survival at
this site was most evident among the 8-mm seed, which
showed declining survival with increased grain size (Table 1).

Experimental Plantings of Mercenaria mercenaria

The shape of the gravel used had no significant effect
(P > 0.05) on clam survival.

TABLE 1.

Experiment I, percent recovery (22 August - 22 October 1980)

of three sizes of seed clams planted in three sizes of gravel

and in sand. Also shown are the total number of mud

crabs (Neopanope sayi) recovered from trays

containing the four substrate types.

Substrate Type
Length of ■ — ■ —

Seed at 6 to 10-mm 10 to 19-mm 19 to 32-mm

Planting Sand Gravel Gravel Gravel Mean

3.9 mm
(n = 4)

7.9 mm

fn = 4)

28.8 mm

14.6

68.4

77.0

1.1
49.6
84.0

0.7
48.6
84.0

0.0

5.8

86.0

4.0
43.1
82.5

Total crabs
recovered 24.0
(n= 10)

36.0

95.0

>306

Final mean shell lengths for the three clam sizes are
given in Table 2. Effects of substrate size or shape on clam
growth were not significant for 29-mm seed (P > 0.05).
High mortality precluded an analysis of growth in the 4-mm
clams. Increasing substrate size did have a significant nega-
tive effect on the growth of 8-mm seed (0.01 >P>0.001).

TABLE 2.

Experiment I, final mean shell lengths (mm) with 95%
confidence intervals (n =12, time = 85 days) for two
sizes of seed clams planted in four types of substrate

Length of
Seed at
Planting

Substrate Type

6 to 10-mm 10 to 19-mm 19 to 32-mm
Gravel Gravel Gravel

Sand

3.9 mm * * *

7.9 mm 15.4 ±2.03 14.0 + 2.21 12.9 ±2.08
28.8 mm 31.8 ± 1.10 33.4 ±3.05 33.0 ±0.12

9.5 ±3.84
31.7 ± 1.48

'Survival was too low to calculate growth rates.

Only a few shell fragments, indicative of crab predation,
were found in the trays containing 4-mm seed. The shells of
these clams were thin enough to be crushed and consumed
by feeding crabs (Landers 1954; Whetstone and Eversole
1978, 1981). Many shell fragments were found in the trays
containing the 8-mm seed. Laboratory studies indicated
that clams of this size can be crushed and consumed by
adult mud crabs, N. sayi (Landers 1954, Whetstone and
Eversole 1978). Shells of dead clams of the larger (29-mm)
seed were primarily paired, intact valves. Several shells had
been cracked, possibly by a large calico crab (O. ocellatus)
or blue crab (C. sapidus). A few shells had chipped or
rasped shell margins suggesting predation by whelks,
Busycon spp. (Carriker 1951, Peterson 1982).

Oyster toadfish (Opsanus tau [Linnaeus] ) were observed
burrowed along the outside edges of three of the trays
throughout the summer and autumn. Three of the four
trays of 4-mm clams planted in sand had survival rates of
3.0, 2.4, and 5.0%. The fourth tray, next to which a toadfish
was burrowed, had a survival rate of 47.3%. Similarly, three
of the four trays of 8-mm clams planted in 10- to 19-mm
gravel contained a mean of seven mud crabs per tray and
had clam survival rates of 48.0, 38.4, and 23.2%. The fourth
tray, which had a toadfish beside it, contained no mud
crabs and had a survival rate of 84.0%. A third toadfish was
found beside a tray containing 29-mm clams. No mud
crabs were found in this tray, but clam survival in that tray
(82%) was not appreciably different from the mean for
clams of that size (82.5%). From these observations, we
hypothesize that the toadfish reduced the abundance of
mud crabs and enhanced the survival of those seed sizes
that were susceptible to mud-crab predation.

Experiment II

Predators observed at Site 1IA were essentially the same
as those listed earlier for nearby Site I. The most abundant
predators observed at Site IIB included calico crabs (Ovalipes
ocellatus) and small knobbed whelks {Busycon carica).
Mud crabs (Neopanope sayi) and small winter flounders
(Pseudopleuronectes americanus) were present but not
abundant.

Significant interactions among the variables tested (size
of seed planted, location, time of planting, and substrate
type) indicated that unqualified general statements about
any single variable cannot be valid (Tables 3 and 4); however,
by considering some of the variables together, some
important results may be noted. All of the variables tested
had significant effects on survival (Table 4). Larger seed
showed better survival than small seed, particularly at Site
IIA. The September-to-June period resulted in better
overall survival than the November-to-August period.
Gravel was generally a better substrate than sand for the
larger clams at Site IIA, but it did not appear to provide
significant survival advantage at Site IIB (Table 3). As in
Experiment I, mud crab colonization was greater in gravel
than in sand.

Experiment III

Dominant predators observed during Experiment III
included small (70- to 80-mm length) knobbed whelks
(Busycon carica), adult (15- to 25-mm carapace width) mud
crabs (Neopanope sayi), and adult (45-mm carapace width)
calico crabs (Ovalipes ocellatus). Abundances of the two
major predator species (B. carica and N. sayi) for which
reliable counts could be made at Sites I IIA, IIIB, and IIIC
are given in Table 5 for three observation dates.

Flagg and Malouf

TABLE 3.

Experiment II, percent recovery (time = 9 months) of three sizes of seed clams in replicate plantings
at two sites in two types of substrate and at two times of the year.

September Planting

November Planting

Site HA

Site IIB

Site HA

Site IIB

Clam Size

Sand

Gravel

Sand

Gravel

Sand

Gravel

Sand

Gravel

3 mm

3.6

0.6

0.0

0.3

0.0

5.7

0.9

0.0

Mean

4.7

0.8

0.0

0.2

0.0

7 mm

6.4

30.4

0.8

4.0

0.0

8.8

0.0

2.4

11.2

16.0

2.4

1.6

0.8

4.8

0.0

Mean

8.8

23.2

1.6

2.8

0.4

6.8

0.0

1.2

23 mm

68.0

94.0

24.0

48.0

50.0

10.0

18.0

68.0

96.0

20.0

42.0

34.0

66.0

26.0

14.0

Mean

68.0

95.0

22.0

45.0

41.0

58.0

18.0

16.0

TABLE 4.

TABLE 5.

Experiment II, four-way analysis of variance (ANOVA) of

percent survival of three sizes of seed clams (3, 6, and

23 mm) planted in two types of substrate (sand and

gravel) at two locations and at two times of the

year (September and November).

Experiment III, abundance of predators (m *) of the two

numerically dominant predator species, the mud crab

Neopanope sayi and the knobbed whelk

Busycon carica.

Source of Variation

Mean Square

d.f.

F Ratio

A = substrate type
B = clam size
C = time of year
D = location

AXB
AXC
AXD
BXC
BXD
CXD
AXBXC

axbxd
aXcxd

BXCXD

aXbxcxd

Within
Total

377.78
7,140.01
1,241.96
2,244.07

187.18

33.60

48.72

148.21

490.22

217.00

75.60

69.87

7.19

4.00

3.93

13.68

1
2
1
1

1
1
2
2
1
2
2
1
2
2

27.61*
521.76*

90.75*
163.99*

13.68*

2.46 n.s.

3.56 n.s.
10.83*
35.82*
15.86*

5.52f

5.11*

0.53 n.s.

0.29 n.s.

0.29 as.

Sampling Date

Neopanope sayi
Site

HIA IIIB IHC

Busycon carica
Site

HIA IIIB IHC

20 May 1981
28 July 1981
14 September 1981

Mean

2.0 0.3 0.0

2.0 0.0 0.0

1.0 0.0 0.0

1.7 0.1 0.0

2.5 1.0 0.3

7.0 2.0 8.6

1.5 1.1 1.2

3.7 1.4 3.4

*significant at 0.01
fsignificant at 0.05
n.s. = not significant
B>D>C>A

In general, survival at Site IIIA was inversely related to
seed size (Table 6). Overall survival was less than 2% even
under the best conditions (3-mm seed in gravel). Only one
of the 6- to 8-mm clams was recovered in July, and by the
termination date (30 September) no clams of that initial

size had survived. No larger seed clams were recovered in
the July sampling. Within a week of planting, empty shells
appeared on the substrate surface.

Maximum recovery cf the 2- to 4-mm seed (in gravel)
was 2.2% at Site IIIB. None of the 6- to 8-mm clams was
recovered, and crushed and cracked shells appeared in the
plots within two weeks of planting. Survival of seed planted
at Site IIIB exceeded 50% only among the 22- to 28-mm seed
clams. Note also in Table 6 that among the 22- to 28-mm
seed there appeared to an initial survival advantage to clams
planted in gravel compared to natural bottom, but by the
time of the final sampling in September, survival rates were
very similar in the two substrate types. Chipped shell
margins and cracked shells indicated predation by whelks
and crabs.

At Site IIIC, survival of the small seed in sand, although
still quite low, was somewhat better than that of the larger
seed sizes (Table 6). By the end of the experiment none of

Experimental Plantings of Mercenaria mercenaria

TABLE 6.

Experiment III, percent recovery of three sizes of seed clams planted at three sites in two types of substrates.

Clams were planted 20 May 1981.

3 mm

7 mm

23 mm

Site

IIIA

llllt

IIIC

Sampling
Date Sand Gravel Sand Gravel Sand Gravel

IIIA

1MB

IIIC

IIIA

IIIB

IIIC

Sand Gravel Sand Gravel Sand Gravel Sand Gravel Sand Gravel Sand Gravel

28 Jul 81
14 Sep 81

3.6
1.5

11.0
1.8

2.0
1.2

3.3

2.2

10.4
6.2

4.5
4.5

0.0
0.0

15.0
0.0

0.0
0.0

62.0
58.0

88.0
61.0

1.7
0.0

14.8
5.3

the 6- to 8-mm clams remained, and a few 23-mm seed
survived only in gravel (5.3%). Heavy losses of the larger
seed clams, the chipped or rasped shell margins of articu-
lated, empty valves remaining in the planting areas, as well
as the high densities of knobbed whelks (B. carica) at this
site (Table 5) suggested that predation by whelks was an
important cause of mortality.

DISCUSSION

The results of this study demonstrated that the character-
istics of a given site, especially the types of predators present,
had an important influence on the loss and presumed
mortality of planted seed clams and on the degree of pro-
tection afforded by recommended culture techniques. For
example, we found that at sites such as Site IIIA where
whelks (Busycon eanaliculatum and B. carica) were
abundant, plantings of 25-mm seed clams suffered complete
mortality despite the presence of gravel aggregate. At
sites such as Site IIIB where mud crabs (Neopanope sayi)
were the dominant predator, the smallest seed clams suffered
high mortality, while the larger seed showed good survival.
Clearly, the idea that seed clams having at least a 25-mm
shell length are relatively immune from most predators
(Menzel 1971, Eldridge et al. 1979) is valid only when the
seed is planted at sites lacking significant populations of
large predators. Existing literature has convincingly shown
that the activity of some important predators such as mud
crabs is significantly reduced by lower autumn temperatures
(Whetstone and Eversole 1981); however, we found that
autumn plantings eventually suffered the same high mortal-
ities as the summer plantings, and the choice of planting
season was inadequate protection against crab predation.

The use of gravel aggregate at Site I, where the mud
crab N. sayi was the dominant predator, gave inconsistent
results in our Experiment I (Table 1). At that site, mortality
among the smaller clams was complete and was independent
of the presence or absence of gravel. On the other hand,
mortality among the larger clams was very low, but it was
again independent of substrate grain size. The survival of
the medium size (7.9-mm) seed was inversely related to
gravel size. The 6- to 10-, 10- to 19-, and 19- to 32-mm

gravels, all of which are within the size range (10 to 30 mm)
used by Castagna and Kraeuter(1977), were not consistently
effective in enhancing seed clam survival (Table 1). Densities
of the mud crab N. sayi were much higher in gravel beds
than in the bare sand (Table 1). There is also evidence from
our data (Table 2) of reduced growth rates among small
seed clams planted in larger gravel compared to those planted
in sand or small gravel.

Gravel may be useful in preventing small clams from
being carried away by currents, although our work offers
no direct evidence for this. It is also possible that gravel and
shell substrates offer more effective protection against
larger crab species than against relatively smaller species
such as N. sayi. Size-related differences in the food and
space utilization of two sympatric xanthid crab species
(Panopeus herbstii and Eurypanopeus depressus [Smith] )
were discussed by McDonald (1982). He noted that the
larger of the two species (P. herbstii) was prevented by its
size from entering narrow spaces between living oysters.
This suggests that the lack of consistent results from seed
plantings in gravel might be due in part to site-specific
differences in the relative abundance of large and small
crabs.

Previous studies have shown that xanthid mud crabs
(primarily N. sayi) are the most abundant clam predators in
Long Island's Great South Bay (MacKenzie 1977). Their
mean, baywide abundance is about 4.4 crabs m~ 2 . while
that of Ovalipes ocellatus is about 0.2 crab m~ 2 (WAPORA,
Inc. 1981). Mud crabs are capable of consuming 1.6 to 5
small (5- to 10-mm) hard clams each day (Landers 1954.
MacKenzie 1977). Theoretically, mud crabs in Great South
Bay could consume up to about 20 seed clams rrf 2 day" 1 .
At this rate of loss, seed plantings of 200 to 500 clams m~ 2
would not survive long. Consequently, local seed planting
efforts that do not somehow protect the young clams
until they are large enough to avoid mud crab predation
will probably be unsuccessful.

Although seed hard clams are readily available from
commercial hatcheries, their cost is relatively high. Costs
for 3- to 5-mm seed range from $10 to $15/1,000 at the
present time (J. Kassner, Town of Brookhaven, NY and

FLAGG AND Malouf

S. Buckner. Town of Islip, NY, pers. comm.). Assuming
that harvested littleneck clams have a dockside value of
about $70 per bag of 500, then the survival and harvest of
planted seed (initially costing $12/1,000) must exceed 9%
of the number planted for the value of the harvest to
exceed the cost of the seed alone. A typical Long Island
town program might plant about 2 million seed and could
require about 6 man-months of handling and planting time.
If the costs of handling and planting are added to the cost
of the seed itself, then the survival requirement might
increase to about 15%. This estimated survival requirement
is relatively low compared to other estimates for commercial
culture (40% by Castagna and Kraeuter 1977, 50% by
Menzel et al. 1976). It should be remembered that our
estimated survival and harvest requirements are minimum
values for seed planted in a public fishery. Existing programs
involve relatively little handling and no maintenance or
protection after planting on the bay bottom. If the costs
of a nursery system (rafts, racks, etc.) were added to our
estimate, the survival requirement for cost effectiveness
would approach those given above for commercial systems.
Our essentially unprotected plantings of 3- to 5-mm seed
clams rarely resulted in survival rates as high as 10%, even in
short-term experiments. Other work, summarized in Table 7,
showed similar results with seed of this size. In fact, 0%

survival was the most commonly encountered result of
unprotected planting of small seed clams. Even when various
types of protective measures were employed, mortality
among small seed clams often exceeded 50% (Table 7).

The relatively low expected survival rates contribute to
the problem of scale in these programs (discussed by
McHugh, 1981). For example, a survival rate of even 15%
would leave only 300,000 clams available for harvest from a
planting of 2 million seed. In the very unlikely event that
all of these clams were harvested, this would yield only
21 m 3 (600 bu), or about 0.6% of each of the three Great
South Bay towns' typical annual harvest. In fact, available
data (Table 7) indicate that survival rates and consequentual
harvest contributions might be much lower.

ACKNOWLEDGMENTS

The authors thank Dr. J. L. McHugh for his critical
review of the manuscript and Charles DeQuillfeldt for his
technical assistance during the study. The cooperation of
Bradden Smith of Shinnecock Tribal Oyster Project, Emil
Usinger of Blue Points Co., Inc., and the East Hampton
Town Council is gratefully acknowledged. Support for this
study was provided by the National Oceanic and Atmos-
pheric Administration, Office of Sea Grant, through the
New York Sea Grant Institute.

TABLE 7.
Published accounts of some trial plantings of seed clams (Mercenaria mercenaria) on the Atlantic coast of the United States.

Reference

Seed Size

Planted

(mm)

Seed Size

Recovered

(mm)

Duration
(Months)

Approximate

Survival

(%)

Notes

Menzel and Sims (1964)

33-44

82-95

33-44

Godwin (1968)

18-22

35-37

18-22

50-52

18-22

36-37

Menzel (1971)

15-35

Walne (1974)

9-13

17-21

Eldridgeet al. (1976)

12-13

16-25

29-45

Menzel et al. (1976)

7-10

0.6

7-10

2.3

7-10

10.1

7-10

58.6

Eldridge et al. (1979)

16-19

46-57

Castagna and Kraeuter (1977)

Kraeuter and Castagna (1977)

1- 3

10-22

Kraeuter and Castagna (1980)

Protection (fence, baited traps)
No protection

No protection

Protection (wire mesh)

Protection (wire mesh; loss due to

"winter-kill")

Protection (fence, traps)

Protection (plastic mesh)

Protection (covered trays)
Protection (covered trays)

No protection
Protection (shell cover)
Protection (gravel)
Protection (wire mesh)

Protection (covered trays)
Protection (same planting as above)

Protection (gravel, traps, baffles)

No protection
Protection (gravel only)
Protection (gravel, baffles)

Protection (pen, gravel, baffles)
Protection (no pen, with gravel, baffles)

EXPERIMENTAL PLANTINGS OF MERCENARIA MERCENARIA

REFERENCES CITED

Carriker, M. R. 1951. Observations on the penetration of tightly
closing bivalves by Busycon and other predators. Ecology
32:73-83.

Castagna, M. 1970. Field experiments testing the use of aggregate
covers to protect juvenile clams. Proc. Natl. Shellfish. Assoc.
60:2 (abstract).

& J. Kraeuter. 1977. Mercenaria culture using stone aggre-
gate for predator protection. Proc. Natl. Shellfish. Assoc. 67:
1-6.

Etdridge, P. J., A. G. Eversole & J. M. Whetstone. 1979. Compara-
tive survival and growth rates of hard clams Mercenaria mercen-
aria, planted in trays subtidally and intertidally at varying
densities in a South Carolina estuary. Proc. Natl. Shellfish.
Assoc. 69:30-39.

Eldridge. P. J., W. Waltz, R. C. Gracy & H. H. Hunt. 1976. Growth
and mortality rates of hatchery seed c\ams,Mercenaria mercenaria,
in protected trays in waters of South Carolina. Proc. Natl. Shell-
fish. Assoc. 66:13-20.

Godwin, W. F. 1968. The growth and survival of planted clams,
Mercenaria mercenaria, on the Georgia coast. Georgia Game Fish
Comm. Mar. Fish. Div. Contrib. Ser. No. 9. 16 p.

Haven, D. & J. D. Andrews. 1957. Survival and growth of Venus
mercenaria, Venus campechiensis, and their hybrids in suspended
trays and on natural bottoms. Proc. Natl. Shellfish. Assoc. 47:
43-49.

Kraeuter. J. N. & M. Castagna. 1977. An analysis of gravel, pens,
crab traps, and current baffles as protection for juvenile hard
clams (Mercenaria mercenaria). Proc. World Maricult. Soc.
8:581-592.

. 1980. Effects of large predators on the field culture of

the hard clam, Mercenaria mercenaria. U.S. Fish Wildl. Serv.
Fish. Bull. 78(2):538-540.

Landers, W. S. 1954. Notes on the predation of the hard clam
Venus mercenaria by the mud crab. Neopanope texana. Ecology
35(3):422.

Mackenzie, C. L. 1977. Predation on hard clam {Mercenaria mercen-

aria) populations. Trans. Am. Fish. Soc. 106(6):530-537.
McDonald, J. 1982. Divergent life history patterns in the co-occurring

intertidal crabs Panopeus herbstii and Eurypanopeus depressus

(Crustacea: Brachyura: Xanthidae). Mar. Ecol. Prog. Ser. 8:

173-180.
McHugh, J. L. 1981. Recent advances in hard clam mariculture.

J. Shellfish. Res. l(l):51-56.
& J. J. C. Ginter. 1978. Fisheries. National Oceanic and

Atmospheric Administration, Marine Ecosystems Analysis

Program (MESA) New York Bight Atlas Monogr. No. 16. 129 p.

Available from: NY Sea Grant Inst., Albany, NY.
MenzeL R. W. 1971. Quahog clams and their possible mariculture.

Proc. World Maricult. Soc. 2:23-36.
, E. W. Cake, M. L. Haines. R. E. Martin & L. A. Olsen.

1976. Clam mariculture in northwest Florida: field study on

predation. Proc. Natl. Shellfish. Assoc. 65:59-62.
MenzeL R. W. & H. W. Sims. 1964. Experimental farming of hard

clams, Mercenaria mercenaria, in Florida. Proc. Natl. Shellfish.

Assoc. 53:103-109.
Peterson, C. H. 1982. Clam predation by whelks {Busycon spp.):

Experimental tests of the importance of prey size, prey density.

and seagrass cover. Mar. Biol. (Berl.) 66:159-170.
Sokal, R. R. & F. J. Rohlf. 1969. Biometry. San Franciso, CA:

W. H. Freeman and Co. 776 p.
Walne, P. R. 1974. Culture of Bivalve Molluscs, 50 Years' Experience

at Conwy. Surrey, England: Fishing News (Books) Ltd. 173 p.
WAPORA, Inc. 1981. Estuarine impact assessment (shellfish

resources) for the Nassau-Suffolk streamflow augmentation

alternatives, draft report on existing conditions. Available from:

U.S. Environ. Protect. Agency, New York. 114 p.
Whetstone, J. M. & A. G. Eversole. 1978. Predation on hard clams,

Mercenaria mercenaria, by mud crabs, Panopeus herbstii. Proc.

Natl. Shellfish. Assoc. 68:42-48.
. 1981. Effects of size and temperature on mud crab,

Panopeus herbstii, predation on hard clams, Mercenaria mercen-
aria. Estuaries 4(2) : 1 53 — 156.

Journal of Shellfish Research, Vol. 3, No. 1, 29-40, 1983.

TRANSPORT OF BIVALVE LARVAE IN JAMES RIVER, VIRGINIA 1

J. D. ANDREWS

Virginia Institute of Marine Science
School of Marine Science
College of William and Mary
Gloucester Point, Virginia 23062

ABSTRACT For nearly 100 years, the James River has been the primary source of seed oysters for Virginia. A disease
caused by Minchinia nelsoni (MSX) killed most oysters in high-salinity waters in the lower river in 1959 and 1960, and
planting has not been resumed in these areas (Andrews 1983). Large populations of oysters on Hampton Bar and near the
mouth of the river which served as broodstocks were destroyed. After 1960, setting declined drastically in regularity and
intensity to about one tenth of that which occurred in the 1950's. Setting patterns suggest two types of seed areas in
Chesapeake Bay: (1) high freshwater discharge, open or flushing estuaries with light spatfalls that decrease in intensity with
distance from the river mouth; the James River is a typical example; and (2) low discharge, trap-type estuaries where
intensive sets are heaviest near the head of the saline sector; examples are the Piankatank and Great Wicomico rivers in
Virginia. Larval transport systems in the two estuarine types differ in quantity of larvae retained and regularity of spatfalls.
Hourly plankton samples in the James River during 10 days in 1964 and 1965 revealed regular cyclic abundance of larvae
with tidal stages. Larvae were 5 to 10 times more numerous during high-tide periods than at low-tide periods. Mostly
early-stage larvae were distributed randomly throughout vertical columns of water. Larvae of other bivalve species exhibited
similar distributions and fluctuations in abundance with tidal stages. Patterns of larval distribution were similar for all
depths at five stations, both in the channel and over oyster beds, during 16 tidal cycles in 1965. Frequent recruitment of
new larval broods and disappearance of most oyster larvae before ages of 3 to 5 days suggest losses due to physical disper-
sion and predation. Only when larvae reached advanced umbo stages did they actively select deeper water strata in the
channel which provided a transport system to carry them upriver. In the 1950's. spatfall occurred every week in the James
River from 1 July to 1 October each year; since 1960, light, erratic setting has prevailed every year. If one assumes that
predation, larval ecology, and physical transport systems have not changed, it appears that broodstocks have become
inadequate, or that larvae were killed by toxic substances.

KEY WORDS: Molluscs, bivalve larvae, transport, distribution, setting (or spatfall), James River, VA

INTRODUCTION

The James River has supplied seed oyster (Crassostrea
virginica [Gmelin] ) for most private grounds in Chesapeake
Bay for over 100 years (Andrews 1951, 1955, 1982a). The
seed area is located in low-salinity waters (< 18 ppt in late
summer) between the James River Bridge and the Deep
Water Shoal (Figure 1). The horizontal salinity gradients
in the James River are steep compared to those of other
estuaries in Chesapeake Bay; salinity in the upper river
seed beds ranges from ppt in late winter and spring
to 10 or 12 ppt in late summer and fall. Consistent annual
spatfalls of moderate intensity averaged 2.7 surviving spat
per shell over 17 years from 1944 to 1960 (Andrews 1982a).
During that period, 90% of surviving spat set on other
oysters. Two to three million bushels (7.0 to 10.6 X 10 4 M 3 )
of seed oysters were harvested annually without depleting
James River stocks. Oysters in the seed area were stunted
in growth and storage of glycogen was low; therefore,
they produced small quantities of spawn; but high-density
populations were spread over large areas of natural shell
beds; no management was applied except for limited
harvesting by hand tongs. Good quality seed oysters with
many single oysters and small clumps resulted from regular

Contribution No. 1180, Virginia Institute of Marine Science

spatfalls and low survival of initial sets (2 to 4% [Andrews
1949] ). Compared to high-salinity areas along the Atlantic
coast of North America, those survival rates were high
(Mackin 1946).

Two types of seed areas are recognized in Chesapeake
Bay based primarily on size of drainage areas and amount
of freshwater discharge (Andrews 1979, 1982b). In the
category of high-freshwater flow are the Susquehanna.
Potomac and James rivers, but only the James permits
recruitment of young oysters with enough regularity and
intensity to be a seed area. Strong freshwater discharge
provides the motive force in these estuaries to establish
strong salinity gradients and a net counterflow of salty
water upriver in the channel; it also produces high flushing
rates to discharge the additional fresh water. The other
category of estuaries, which I call trap-type seed areas
(Andrews 1979), consists of low-discharge rivers with
small drainage areas. Two examples of this type seed area
which have been studied are the St. Marys River (Manning
and Whaley 1954) for distribution and retention of larvae,
and the Manokin River (Carter 1967) for circulation
regimes. Other important seed areas in Chesapeake Bay
which belong in this trap-type category are the Piankatank
and Great Wicomico rivers in Virginia, and Broad Creek, a
branch of the Choptank River in Maryland (Boicourt 1982).

ANDREWS

OLD
CHANNEL

BURWELL
BAY

Figure 1. Map of James River seed area from Hampton Roads to last upriver seed bed at Deep Water Shoal. Sampling stations
and associated oyster beds are designated in kilometers from mouth of the river.

Transport of bivalve Larvae

The oyster setting patterns in these high-flushing and
trap-type estuaries reflect differences in circulation patterns
that result in dispersion or retention of larvae. The James
River is the only flushing-type estuary in Chesapeake Bay
with adequate spatfall to be a seed area. Spatfall was con-
sistent annually, but from low to moderate in intensity; it
exhibited a gradient of declining setting intensity from the
mouth to upriver areas (Andrews 1982a). The gradient of
setting was reversed in trap-type estuaries with highest
spatfalls on the upriver beds (Manning and Whaley 1954,
Andrew's data in Haven et al. 1978). For comparison,
setting was consistent in intensity and regular by years in
the James River; but intensity was much higher in trap-type
estuaries and quite irregular by years with frequent failures.
There was no change in the patterns of spatfall in trap-type
estuaries following introduction of the disease caused by
Minchinia nehoni (MSX) to Chesapeake Bay in 1959
(Andrews and Wood 1967); but in the James River there
was a severe reduction in setting intensity and spatfall
became erratic in distribution (Haven et al. 1978). All
seed areas in Chesapeake Bay are in low-salinity (< 20 ppt)
waters and usually not subject to MSX infections and
mortalities; broodstocks were greatly reduced in the lower
James River by MSX, but they were not in the trap-type
seed areas which are located upbay and lay mostly above
the endemic area for the disease.

The geography and morphology of the two types of
estuaries are probably significant factors with respect to
dispersion and retention of larvae (Andrews 1979). The
James River has a wide, deep channel, bordered by wide,
shallow flats where oyster beds are located; it has few
tributaries and limited marsh areas adjacent to the oyster-
growing sector. The trap-type seed areas have meandering
channels, numerous projecting points, very shallow flats,
and many tributary creeks. Reduction and deflection of
currents by boundary effects and morphometry in these
tortuous estuaries probably aid in retention of larvae. The
Great Wicomico River is an excellent example of the
morphology of a trap-type estuary with its characteristics
of infrequent but intensive spatfalls. Over 30 years, failures
have been more frequent than successes in the Virginia
trap-type rivers (Haven et al. 1978).

The first study of larval transport in Chesapeake Bay was
conducted in the James River in 1950 by the Virginia
Fisheries Laboratory and the Chesapeake Bay Institute (CB1)
(Pritchard 1953). An intensive study of physical and
chemical hydrology was conducted by CBI (Pritchard
1952, 1955). Concurrently, bivalve larvae were sampled
bi-hourly by Virginia biologists at three stations across the
river at the Wreck Shoal (J 17) level (Andrews 1982c).
Wreck Shoal is the largest and most productive oyster bed
in the James River. The last period of sampling, from
30 August to 3 September, coincided with peak setting of
oysters in that year with 40 spat per shellface per week on
four replicate shell strings that were suspended off the

bottom at Wreck Shoal (Andrews 195 1 ). Larvae were scarce
at all stations and all sampling depths (3 depths in channel,

2 over beds). Primarily, straight-hinge larvae of less than

3 days of age were found, and many samples had no oyster
larvae. Advanced larvae were encountered only rarely even
when volume of plankton samples was increased from 100
to 500 C (Andrews 1982c). Preliminary data on larval
densities were presented by Pritchard (1953) who calculated
that only one mature larva per 100 C was needed to produce
the observed spatfall. No conclusions were reached about
distribution systems for larvae and for their retention in the
seed area.

The studies of Manning and Whaley (1954) in St. Marys
River, Maryland, a trap-type estuary, were far more conclu-
sive because advanced larvae were abundant and they
moved upriver with wind-induced currents. Larvae in all
stages were found and often 100 or more late-umbo larvae
in 100-C samples. Densities of advanced stage larvae were
much higher in deeper waters in the channel with peak
counts of 900 late-umbo larvae per 100-2 sample. Manning
and Whaley concluded that wind-induced convection
currents moved surface waters landward in the lower-river
sector with downriver flow in bottom layers. The typical
characteristics of trap-type seed areas with tortuous geog-
raphy and most intensive spatfalls near the head of the
estuary are illustrated in Figure 1 of Manning and Whaley
(1954).

Carter (1967) conducted a physical study of hydrography
of Manokin River on the Eastern Shore of Maryland using
point release of dye to simulate physical dispersal of
larvae. His conclusions were similar to those of Manning
and Whaley (1954) that wind-induced convection currents
carried larvae upstream. Freshwater discharge was almost
negligible as in St. Marys River. Although the Manokin
River is not a seed area, it could be according to Carter if
enough brood oysters were planted in the lower river.
Seliger and Boggs (1983 ) examined the physical hydrography
of the Choptank River and its tributaries; they confirmed
the physical regimes of trap-type estuaries but provided
little information on larval biology from limited sampling,
except that larvae were most abundant at the heads of
saline river systems (creeks) where setting is known to be
highest (Meritt 1977). More detailed studies of circulation
in tributary creeks of the Choptank River were made by
Boicourt (1982).

Mechanisms of transport and setting of planktonic larvae
in other estuaries are discussed by Ketchum (1954) in
general, by Korringa (1952) for oysters in the Oosterschelde
(Holland), and by Carriker (1951), Nelson (1957) and
Haskin (1964) for oysters in New Jersey coastal bays and
Delaware Bay. There is considerable literature on upstream
movements of fish and crustaceans (e.g., Sulkin 1981), but
larvae and juveniles of these groups make more positive
responses to favorable strata and currents than do bivalve
larvae. The most important bivalve larval studies of open

ANDREWS

systems such as James River are those of Kunkle(1958) and
Hidu and Haskin (1971) along the Cape May shore in
Delaware Bay. In 1964—1965, mature and eyed-larvae were
abundant in 200-C samples collected by the latter authors
with 160-/im mesh plankton nets, and setting was intense.
This area consistently had intense spatfalls (Nelson 1959),
often far higher than any place in Chesapeake Bay. Delaware
Bay is similar to James River in physical characteristics,
but it has lower freshwater discharge than does Chesapeake
Bay (Boicourt 1982). It has a tidal range of nearly 2 m,
which is twice that of Chesapeake Bay (x = 0.72 m). Tidal-
and wind-induced mixing in this wide, shallow bay, as in the
James River, prevent much vertical density stratification
in summer. By Pritchard's (1955) criteria for circulation
regimes, both estuaries are type C in summer with lateral
mixing; because of decreased river discharge and wide,
shallow basins, salt balance is maintained by circular flow
(Pritchard 1956).

This report describes the patterns of larval transport in
the James River and compares transport of larvae in the
two types of estuaries. During 22 years (1946 to 1967) of
intensive monitoring of spatfall in James River, the final
distributions of larvae were determined (Andrews 1951,
1955, 1982a), but how they became distributed throughout
the seed area is still obscure. The importance of large
broodstock populations was shown after 1960, when setting
rates declined to less than one-tenth the 1950's level;
this followed cessation of private oyster planting in the
lower river (Haven et al 1978, Andrews 1982a). High
mortalities caused by MSX prohibited use of James River
seed oysters in high-salinity waters of the lower river
(Andrews 1983). Scarcity of oyster larvae during the 1960's,
particularly of advanced stages, made studies of larval
ecology difficult. Descriptions of the two types of seed
areas are based primarily on patterns of spatfall that
indicated wide differences in retention of larvae. Larval
studies have not been made in trap-type estuaries in Virginia.
Dye studies conducted in a physical model of James River
at Vicksburg, Mississippi, suggested the probable extent of
larval dispersion if transport were passive (Hargis 1966).
Only field data collected in James River when sampling was
most intensive in 1964 and 1965 are reported here. Data
for earlier larval studies in James River are reported by
Andrews (1982c). Some physical data collected during the
8 days of plankton samplingin the 1965 study were reported
by Wood and Hargis (1971 ).

MATERIALS AND METHODS

Scarcity of larvae at Wreck Shoal in 1950 and recognition
of higher spatfalls in the lower river resulted in selection of
the Brown Shoal area for sampling in 1964 and 1965. Based
on intensity of spatfalls over 20 years and preliminary
plankton samples each year, a period near 1 September was
chosen as the optimum time for sampling. This would not
be true of any other estuary in Chesapeake Bay because the

James River always has late setting. More emphasis was
placed on sampling in the channel than over inshore oyster
beds because deep-water currents are necessary for physical
transport upriver. The channel is considered to be the
primary transport route for upstream movement of larvae.
Sampling was conducted hourly during night and day at
four depths (0, 3, 6, 9 m) in the channel and at two depths
over 3-m-deep beds for 2 days in 1964 and 8 days in 1965.
After finding early-umbo larvae in the channel at Brown
Shoal on 31 August 1964, stations were established at J33
in the channel and at Wreck Shoal (J33E) bed where
sampling occurred for one tidal cycle on 3 September 1964.

Three vessels were spaced 2 km apart and anchored in
the channel in 1965, and two were anchored inshore over
oyster beds opposite the central channel station above the
James River Bridge. All plankton samples were taken
synoptically on the hour with submerged pumps for each
depth. Volume of water was measured by timing of calibrated
pumps. Samples of about 300 C were pumped into plankton
nets with 50-nm mesh submerged in watertight boxes.
Surface and bottom samples were taken 1 m from interfaces
with air and substrate to avoid boundary effects on currents
and larvae.

Plankton samples were preserved with 1% formalin
buffered with an excess of NaHC0 3 or NaBr0 3 crystals.
Counts of all species of bivalve larvae were made on
Sedgwick-Rafter cells. In 1964, three or more 2-cm 3
aliquots were pipetted from magnetically stirred samples
condensed to about 60 cm 3 . In 1965, entire samples were
counted after excess fluid was decanted; sediments were
swirled in 10-cm watch crystals to remove lighter periferal
plankton and fecal pellets with pipettes. Several slides
were counted for each swirl depending on the amount of
sand and sediment; three or more swirls were made for
each sample until larval counts declined rapidly. Early-
stage larvae are lighter than advanced larvae, therefore they
are more difficult to separate from other plankton by this
swirling method. Total sample counts were necessary
because of low density of larvae. All species were counted
separately by stages of development; these were designated
as straight-hinge, early-umbo, late-umbo, and mature or
setting-size larvae (Chanley and Andrews 1971). Species
and stages with low abundance were not summarized except
as total bivalve larvae. Oysters comprised about one half of
the bivalve larvae in most samples.

RESULTS

Brown Shoals was sampled hourly through one tidal
cycle on 31 August 1964. A density of 10 to 40/8 of
early-stage oyster larvae with some advanced larvae was
encountered. A severe thunderstorm interrupted this field
study at midnight, but a new operation during one daytime
tidal cycle was carried out at J 19 and J33 on 3 September
1964. Counts of total bivalve larvae in the channel at J19
are shown in Table 1. Bivalve larvae were two to several

Transport of Bivalve Larvae

times more abundant at 3- and 6-m depths than at and 9 m
near surface and bottom boundaries. Larvae at 3 m depth
had reached abundances of 30/2 at maximum flood tide
and stayed high through high-slack water to maximum ebb.
It is clear, however, that larvae were patchy in local distri-
bution at various sampling times. A new group of early-
stage larvae, 2 to 3 days old, had entered the Brown Shoal
area on 3 September, and advanced larvae were less abun-
dant than they had been on 3 1 August.

TABLE 1.

Total of bivalve larvae per 10 liters by depths in channel

at Brown Shoal (J 19), James River,

3 September 1964*

Bivalve Larvae by Depth (m)

Time

Tide

1000-1100

early flood

—

1100-1200

118

228

158

1200-1300

387

676

278

1300-1400

maximum flood

298

118

1400-1500

529

163

1500-1600

483

170

1600-1700

high slack

424

397

1700-1800

111

341

263

124

1800-1900

maximum ebb

189

640

222

168

Mean

328

233

105

*70% oyster larvae

Samples at station J33 in the Wreck Shoal area on
3 September 1964 showed that advanced oyster larvae had
moved upriver (Table 2). This table is arranged to show
increasing densities of advanced-stage larvae with greater
depths. Advanced larvae were much less abundant inshore
over Wreck Shoal at station J33E in 3 m of water than in
the channel. Again, patchiness of larvae was evident although
some late-umbo larvae were found at all depths sampled.
These counts were made by P. Chanley and the first 50
larvae were measured for size. This was the only one of
17 days sampled during full-tidal cycles over four years
(1950, 1963, 1964, 1965) when significant numbers of
advanced oyster larvae were found in James River. A light
spatfall from these larvae occurred throughout the seed area
in two subsequent weeks (Andrews 1982a).

Hourly sampling around the clock from 5 and 3 stationary
vessels, respectively, for 8 days (30 August to 3 September
and 9 to 1 1 September) in 1965 showed bivalve larvae in
regular cycles of abundance with tidal stages. High abun-
dances occurred from maximal flood velocities through
high-slack water to maximal ebb velocities, and low densities
occurred during the other half of each tidal cycle. Combined
totals for all bivalve larvae for four depths in the channel
are shown for two stations (Figure 2). Most larvae of all
species, including oyster larvae, were at straight-hinge
stage (Andrews 1982c). Data for total bivalve larvae by four
depths at one channel station exhibited similar patterns
of cyclic abundance (Figure 3). Early-stage larvae were

TABLE 2.
Population densities of advanced oyster larvae (number per liter) by depths in channel at Wreck Shoal (J33), 3 September 1964.

Oyster Larvae by Depths (m) and by Sizes (pm) 1

3.5-4.0

Time < 125 125-200 >200 < 125 125-200 >200

<125

7.0-8.0

125-200

>200

1125

1208

1227

1300

349

1325

1345

1359

429

1420

1442

1500

285

1522

1544

1600

177

1624

1646

1701

406

1725

1743

1800

202

Mean

266

1877
818

698

550
166
318

63
160
128

738

412

201

427

190

252

218

197

138

137

215

103

Stages of larvae by size are: straight-hinge = < 125 /im; early-umbo = 125 to 200 /Jm; late-umbo or eyed = > 200 )Jm.

ANDREWS

TIDAL
VELOCITY

IOOO 1400 1800 2200 0200 0600 1000 1400 1800 2200 0200 0600 1000

September 9

September 10

September 11

Figure 2. Hourly densities of total bivalve larvae at four combined depths in channel, 9 to 11 September 1965. Two sampling stations
designated by anchored vessels R/V LANGLEY and R/V PATHFINDER in channel 2 km apart. Total counts from 300-? samples at four
depths adjusted to number per 100 5. Similar cycles of abundance occurred each tidal cycle at five stations over a period of 8 days between
30 August and 1 1 September 1965. Early-stage larvae predominated throughout the period.

o
o

300

a- 200

<r>

2
3

100

I METER DEPTH

4 METER DEPTH

7 METER DEPTH

10 METER DEPTH

~i r^ — I 1 1 1 1 1 1 1 1 r

1000 1400 1800 2200 0200 0600 1000

9 SEP

1 1 1 1 1 I 1 1 1 1

400 1800 2200 0200 0600 1000

10 SEP

II SEP

Figure 3. Cyclic abundance of bivalve larvae with tidal stage by depths in channel. Samples taken simultaneously with four submerged
pumps at four depths at station J 19.

TRANSPORT OK BIVALVE LARVAE

distributed throughout vertical columns of water with
highest densities usually at 4 and 7 m.

Data on bivalve larvae by species also showed highest
densities from mid-flood to mid-ebb tidal velocities
(Figure 4). Patchiness was evident, but peaks of abundance
for oysters and other bivalves tended to occur near high-
slack-water stage. Highest densities at high tides were 5 to
10 times as great as lowest densities at low tides. Oyster
larvae were the most abundant of bivalve species, but peak
densities tended to occur concurrently for all species.

The cyclic abundance of larvae in shallow waters (< 3 m)
over oyster beds is illustrated in Figure 5. High and low
densities appeared at the same tidal stages as in the channel
but tended to differ more widely in densities.

DISCUSSION

Oyster spawn is released at least weekly during summer
from late June through September in the James River, but
spatfall is most successful in late August and early September
(Andrews 1955). Although spatfall occurred every week
from 1 July to 1 October in the 1950's, 25 years of setting
records indicate that conditions for survival and transport
of larvae are most favorable in late summer (Andrews
1982a). This is a period of low-freshwater discharge and
high salinities; therefore, stratification is minimal and net
upriver movement of saline water in the channel at depths
below 3 m is small and slow (Pritchard 1953, 1955).
Nevertheless, in contrast to trap-type estuaries, the James
River always has freshwater discharge which induces some
stratification and mixing upriver in the seed area. Hampton

Roads is nearly homogeneous for density of water in late
summer, yet some saline water must move upstream in the
channel to maintain salt balance in the seed area. Salinities
increase gradually in the seed area as summer progresses.

Dye releases near the mouth of the James River in the
Vicksburg model showed that a 28.3-m 3 /s ( 1,000- ft 3 /s) dis-
charge rate, which approximated salinity regimes observed
in late summer of 1964 and 1965, resulted in higher concen-
trations of dye at Burrells Bay after seven prototype days
than a 90-m 3 /s (3,200-ft 3 /s) discharge (Hargis 1966). This
suggests less importance of salt-balance transport upriver
and greater effects of high-flushing rates that remove larvae
from the river. If tidal dispersion is the primary factor or
transport system regulating distribution of bivalve larvae,
late-summer hydrographic regimes would be most favorable
for retention of larvae in the river.

Oyster larvae originate over shallow inshore flats and
oyster beds in the James River. Early-stage larvae occur in
the full vertical column of water over flats and in the
channel; therefore, most larvae released in the seed area are
probably carried downriver in shallow surface waters during
their first days of planktonic life. Before MSX stopped the
planting of seed oysters in Hampton Roads, a large oyster
population near the river mouth supplied large quantities of
spawn. In post-MSX years after 1960, most larvae originated
in the seed area. The topography of the river below the
James River Bridge delivers larvae off the extensive eastern
shore seed beds into the channel of Hampton Roads where
a deep-water column of 10 m or more is thoroughly mixed
and available to allow vertical redistribution of larvae for

O
O

UJ
Q.

<
>

200

w 100-

>
_j
<

Crassostreo
Mull ma
Anomia
Other Bivalves

1000

"i — i — i — i — f -\ — i — i 1 — i r — i — i — i — i

1400 1800 2200 0200 0600 1000 1400 1800 2200 0200 0600 1000

9 SEP 10 SEP I I SEP

Figure 4. Cyclic abundance of bivalve larvae by species. Highest densities occurred between maximal flood and maximal ebb stages of tides.

ANDREWS

k 300-

CC
UJ

>
a.
<

UJ
CD

2
3

200

100

1000

1800
9 SEP

2200 0200 0600

n r

1000

1400
10 SEP

i 1 1 1 r

1800 2200 0200

0600 1000
I I SEP.

Figure 5. Density of bivalve larvae at surface and bottom over Brown Shoal oyster bed. Abundance of larvae was lower over shoals but
cyclic patterns with tidal stages were similar for species and depths.

river ascent in the channel. Early-stage larvae appear to be
recycled several times up the channel, out over the flats,
and back down to Hampton Roads during their first days of
pelagic life. Most larvae disappeared within less than 5 days;
they were replaced by newly spawned larvae. Few larvae
achieved advanced umbo stages during which they would
have selected deeper layers of water thereby enabling them
to ascend into the seed area.

My data and concept of transport and dispersal of bivalve
larvae apply primarily to early-stage larvae (Figure 6). The
seed area provides the larvae and Hampton Roads is a deep-
mixing zone which facilitates advection of larvae upriver
in the channel. These are primary but not exclusive roles
for the two river sectors shown in the diagram. It is apparent
from plankton sampling and spatfall patterns that new
groups of young larvae are being introduced every week, or
more frequently. Larvae in waters discharged into Chesa-
peake Bay are lost at an estimated flushing rate of 15% per
tidal cycle (A. Kuo, Virginia Institute of Marine Science,
Gloucester Point, VA; pers. coram.); this sums to 95% loss

of larvae in 10 days or 20 tidal cycles, the shortest probable
duration of larval life in nature. Data on larval abundance
near the river mouth are not available, but it is presumed
from the spatfall gradients that eventually setting-size larvae
are at least as abundant as at Brown Shoals. Hourly sampling
during 5- and 3-day physical and biological studies in a 13-
day period in September 1965 showed the scarcity of
advanced oyster larvae in the James River. Larvae were not
surviving in the James River long enough to grow to umbo
larvae (3 to 5 days) and, therefore, could not utilize the net
upriver channel flow in waters greater than 3 m depth.
There are no data on losses of bivalve larvae by predation in
nature, although my assumption is that the same predators
present in the 1950's are still equally active in the 1960's
and 1970's. Many pelagic larvae, including fish fry, coelen-
terates, ctenophores, as well as most adult bottom-living
organisms with mucus and ciliary feeding mechanisms,
capture bivalve larvae (Mileikovsky 1974, Andrews 1979).
Most efficient as collectors are adult oysters on beds where
mature larvae are most attracted by gregarious setting.

Transport of Bivalve Larvae

TRANSPORT OF BIVALVE LARVAE IN THE JAMES RIVER

FRESHWATER DISCHARGE

FLATS

MID-FLOOD I

VESSEL STATIONS 1966

OR MID-EBB -V"

MID- EBB /

TO _l

MID- FLOOD,

POSITION OF

MAJOR LARVAL

BROODS

JAMES RIVER BRIDGE

TIDAL CURRENTS

SALINE WATER INPUT

SEED OYSTER
BEDS (SPAWNING)

SEED

OYSTER

AREA

HAMPTON
ROADS

CHESAPEAKE
BAY

LARVAE
LOST

Figure 6. Diagram of a hypothesis of larva] transport in James River. Oyster beds and larval broods are located only symbolically. Channel
transport is emphasized, but transport of larvae occurs throughout cross sections of the river. Width of arrows suggests intensity of transport
system and density of larvae. A tidal excursion is about 1 1 km in channel. The bridge and Deep Water Shoal are 19 and 46 km, respectively,
above the river mouth.

ANDREWS

Figure 6 emphasizes the importance of channel waters
for transport of larvae upriver. Tidal excursions average
about 1 1 km in the channel; this means that larvae located
at the bridge could be carried to Wreck Shoal in one flood
tide, or downriver to the middle of Hampton Roads in one
ebb tide. In three years (1963—1965) of late-summer
sampling in the Brown Shoal area, oyster larvae were rarely
absent; this indicates that one or more broods were dis-
tributed at least 1 1 km above and below the bridge during a
tidal cycle. The larval groups illustrated by ovals on Figure 6
are intended to suggest the location where larvae were most
abundant at given tidal stages. The arrows suggest densities
of larvae in the channel and at sites of dispersion over
oyster beds. Most larvae carried upriver during flood tide
appear to be carried back down the channel during ebb
tide; a few must be trapped over shallow oyster beds or in
meandering creeks by eddies and boundary effects (slowing
of currents) of bottom and marginal features such as
marshes. Apparently, advanced larvae at Wreck Shoal on
3 September 1964, which were abundant mostly in the
channel, reached oyster beds in the seed area by slow
advance in net upstream flow in deep channel currents.

Wood and Hargis (1971) reported on a 24-hour period
of sampling(l September 1965) during the same field study
reported in this paper. Larvae showed the same patterns of
abundance given in this report and also in the other days
not reported by either of us. In their samples, oyster larvae
were usually fewer than 100 per 300-8 sample, although
early-umbo-stage larvae were relatively abundant. They
reported physical data on circulation, salinity, temperature,
and net flow based on seven complete tidal cycles of
observation. These physical conditions apply equally well to
plankton data presented in this paper for 9 to 1 1 September.
The type C counter-clockwise circulatory pattern described
by Pritchard (1955) prevails in the James River in late
summer when freshwater discharge is low. Monthly river
discharge averaged less than 28.3 m 3 /s (< 1,000 ft 3 /s) for
the months of August and September 1964 and 1965. Net
upriver flows are greater on the northeastern side of the
channel, and discharge is greatest downriver on the south-
western shore.

Wood and Hargis (1971) contended that oyster larvae on
the bottom responded to salinity stimulation during flood
tides, but they provided no data that showed selective
swimming or distribution of larvae by depths. Vertical
salinity gradients in Hampton Roads where larvae originate
with each flood tide were less than 1 ppt from surface to
bottom. If larvae rested on the bottom during ebb and low
tides, they could respond to increasing salinities during
flood tides (Haskin 1964), but evidence that larvae rest on
the bottom is inconclusive. Carriker (195 1) worked in high-
salinity coastal bays where shallow water and strong pycno-
clines prevented larvae from freely selecting strata for
upriver transport. Both Carriker (1950) and Wood and
Hargis (1971) support Nelson's hypothesis (Nelson and

Perkins 1931) that oyster larvae ascend estuaries by resting
on the bottom during ebb tides and by swimming during
flood tides. Data of Wood and Hargis (1971) comparing
coal particles with larvae seem irrelevant to me because it
has been clearly established that bivalve larvae can move
vertically by their own powers of swimming. Larvae were
found during all tidal stages whereas coal particles were
observed only during strong currents. Larvae were most
often abundant at high-slack water and there was no
evidence that larvae descended during periods of slack
currents. Larvae were least abundant in samples taken near
the bottom during strong tidal currents when large numbers
of fecal pellets (primarily from oysters) and sand grains
were found in samples. This leads me to believe that larvae
are actually trapped on the bottom during strong currents
by the roiling effects of bottom drag and constant pelting-
even though all are being carried by slow bottom currents.
Dirty samples taken too close to the bottom always con-
tained few larvae. If distribution of larvae were completely
passive, they would spend both high- and low-slack periods
on the bottom just as coal particles and fecal pellets do,
but feeding time would be reduced. Losses of larvae to
smothering and predation on the bottom may be as great
as those from dispersal and predation during planktonic
life.

Counts of larvae collected through 8 days (16 tidal cycles)
show that the pattern of highest abundance from mid-flood
to mid-ebb tides was regular and highly significant, but
explanations of cyclic abundance vary in the literature. The
important observations of the present study are: (1) total
quantities of larvae at all stations before and after slack-high
water were approximately equal; (2) persistence of early-
stage larvae indicated that new broods were recruited fre-
quently into the river; (3) older larvae were found most
frequently in deeper waters and, therefore, in the channel;
and (4) there was a noticeable decrease in density of larvae
from the lower channel station to the upper one, only 4 km
apart, at all tidal stages.

Larval broods are three dimensional. The term swarm is
inappropriate for there is no evidence that larvae remain
together or aggregate horizontally. Advanced larvae choose
deeper strata in the water column effectively. Passive
physical transport probably far outweighs in significance
any results from selective motion by larvae, particularly
during the first 5 days of planktonic life. Larvae do respond
to pheromones when setting is about to occur. It is not
known whether they can respond to food or other stimuli.

My scenario for the decline of setting in James River
since 1960 assumes that loss of brood stocks to MSX disease
in the lower river resulted in too few larvae to replenish
oyster stocks in the seed area. It appears that broods of
larvae are carried up and down the river several times with
progressive thinning and dispersal of each brood. In the
area sampled in 1965, near the James River Bridge, larvae
probably moved up the channel and along the northeastern

Transport of Bivalve Larvae

shallow flats, then back down the channel and over the
southwestern flats to Hampton Roads (Wood and Hargis
1971). Most larvae were lost by dispersion and predation in
3 to 5 days before they were stimulated to swim in deeper
strata. New broods replaced old ones repeatedly. Spring
tides and storms that increase tidal amplitude over the
mean 0.72 m may cause some larvae to be trapped inshore
and result in spatfalls. Because the same circulatory patterns
still exist in James River, regular spatfalls every week for
3 months in the 1950's may be attributed to much larger
populations of brood oysters and greater abundance of
larvae in that period.

In the mid-1960's, Langley Wood (VIMS, Gloucester
Point, Virginia, unpublished studies) constructed a vertical
plexiglass cylinder about 2.5 m long and 0.3 m in diameter
to study the swimming habits of oyster larvae. A strong light
was mounted over the upper end and sampling ports were
inserted at various levels. Larvae alternated between
swimming upward in gyrals and falling slowly while resting
for periods of a minute or so. When larvae bumped into one
another they quickly retracted their velums. Pelagic larvae
have two purposes: to distribute the species and to replenish
adult stages (Galtsoff 1964). The velum provides a mechan-
ism for swimming and feeding activities to meet these
goals. Larvae must swim to eat. Resting for half of each
tidal cycle on the bottom may require a doubling of the
duration of larval life. In hatchery cultures, strong light
causes swimming larvae to seek shade and curious distri-
butional patterns visible to the naked eye are formed. In
many estuaries, larvae are confronted with unfavorable
natural conditions such as low temperatures or toxic com-
pounds below surface waters (Quayle 1969). In these waters
larvae are forced to swim continuously throughout their
planktonic life regardless of dispersal effects.

I conclude that bivalve larvae swim continuously during
larval life and that their dispersal and ultimate fates are
strongly dependent on current regimes and flushing rates of
estuaries. The bottom is a hazardous place for larvae to
rest: a host of sedentary filter feeders become predators or
imprison larvae in mucous-wrapped fecal pellets (Cerruti
1941,Mileikovsky 1974). Siltation is a serious threat on the
bottom in channels where currents are strong. Prolonged
duration of larval life and exposure to predators are major
threats to survival in the James River with its relatively
high flushing rates. The trap-type estuaries with their rela-
tively intensive setting rates provide physical transport
regimes that allow greater retention of larvae. If oyster
larvae can persist in an estuary long enough to reach umbo
size, a preference for deeper waters prevails and, in the case
of the James River, they should be able to ascend the
deep channel currents more effectively than in the poorly
stratified trap-type estuaries. Observations from setting
records indicate that the opposite occurs and that they are
less successful in remaining in strong flushing-type estuaries.
This implies that passive physical transport predominates
over larval reactions to physical and chemical stimuli to
select favorable current strata. Presumably, more intensive
oyster setting in Delaware Bay can be attributed to the
large size of the estuary with lower freshwater-discharge
rates and to its wide shallow flats; only the upper seed area
sector exhibits type-C circulation in summer, and flushing
rates in the widened lower sector (Hidu and Haskin 1971)
are probably much lower than in James River.

ACKNOWLEDGMENTS

I acknowledge the dedicated support of Martha Eble.
Sybil Lawler, Paul Chanley. Donna DeMoranville, and Ed
Powell who counted larvae in many plankton samples
during 1965 and 1966.

REFERENCES CITED

Andrews, J. D. 1949. The 1947 oyster strike in the James River.

Proc. Natl. Shellfish. Assoc. (1948):61-66.
. 1951. Seasonal patterns of oyster setting in the James

River and Chesapeake Bay. Ecology 32:752-758.
. 1955. Setting of oyster in Virginia. Proc. Natl. Shellfish.

Assoc. 45:38-46.
. 1979. Pelecypoda: Ostreidae. Giese, A. C. and J. S. Pearse,

eds. Reproduction of Marine Invertebrates. Vol. 5. Molluscs:

Pelecypoda and Lesser Classes. New York, NY: Academic Press.

.198 2a. The James River public seed oyster area in Virginia. Va.

Inst. Mar. Sci. Spec. Sci. Rep. Appl. Mar. Sci. OceanEng. 26 1 : 60 p.
. 1982b. Reproduction of oysters in Virginia. Available

from author on request: Virginia Institute of Marine Science,
Gloucester Point, VA. (unpublished manuscript)
. 1982c. Transport of the bivalve larvae in the James

River, Virginia. Va. Inst. Mar. Sci. Spec. Sci. Rep. 1 1 1 : 75 p.
. 1983. Minchinia nelsoni (MSX) infections of oysters in

the James River seed area and their expulsion in spring. Estuarine
Coastal Shelf Sci. 16:255-269.
& J. L. Wood. 1967. Oyster mortality studies in Virginia.

VI. History and distribution of Minchinia nelsoni, a pathogen of

oysters in Virginia. Chesapeake Sci. 8:1-13.
Boicourt, W. C. 1982. Estuarine larval retention mechnisms on two

scales. Kennedy, V. S., ed. Estuarine Comparisons. New York,

NY: Academic Press, p. 445-457.
Carriker, M. R. 1951. Ecological observations on the distribution of

oyster larvae in New Jersey estuaries. Ecol. Monogr. 21 : 19—38.
Carter, H. H. 1967. A method for predicting broodstock require-
ments for oyster (C. virginica) producing areas with application

to the Manokin River. Chesapeake Bay Inst. Johns Hopkins Univ.

Spec. Rep. 13: 37 p.
Cerutti, A. 1941. Osservazioni ed esperimenti sulle cause di distru-

zione delle larve d'ostrica nel Mar Piccole e nel Mar grande di

Taranto. Arch. Oceanogr. Limno. 1:165-201.
Chanley, P. & J. D. Andrews. 1971. Aids for identification of bivalve

larvae of Virginia. Malacologia 11:45-119.
Galtsoff, P. S. 1964. The American oyster Crassostrea virginica

Gmelin. U.S. Fish Wildl. Serv. Fish. Bull. 64: 480 p.
Hargis, W. J., Jr. 1966. Operation James River, an evaluation of

physical and biological effects of the proposed James River

navigation project. Va. Inst. Mar. Sci. Spec. Sci. Rep. Appl. Mar.

Sci. Ocean Eng. 7: 73 p.

ANDREWS

Haskin, H. H. 1964. The distribution of oyster larvae in Delaware

Bay. Nanagansett, RI: Proc. Symp. Exp. Mar. Ecol., Occas.

Publ. 2:76-80.
Haven, D. S.. W. J. Hargis, Jr. & P. C. Kendall. 1978. The oyster

industry of Virginia: its status, problems and promise. Va. Inst.

Mar. Sci. Spec. Pap. Mar. Sci. 4: 1024 p.
Hidu, H. & H. H. Haskin. 1971. Setting of the American oysters

related to environmental factors and larval behavior. Proc. Natl.

Shellfish. Assoc. 61:35-50.
Ketchum, B. H. 1954. Relation between circulation and planktonic

populations in estuaries. Ecology 35:191-200.
Korringa. P. 1952. Recent advances in oyster biology. Q. Rev. Biol.

27:266-308,339-365.
Kunkle, D. C. 1958. The vertical distribution of oyster larvae in

Delaware Bay.Proc. Natl. Shellfish. Assoc. 48:90-91.
Mackin, J. G. 1946. A study of oyster strike on the Seaside of

Virginia. Va. Fish. Lab. Contr. No. 25: 18 p.
Manning, J. H. & H. H. Whaley. 1954. Distribution of oyster larvae

and spat in relation to some environmental factors in a tidal

estuary. Proc. Natl. Shellfish. Assoc. 45:56-65.
Meritt, D. W. 1977. Oyster spat set on natural cultch in the Maryland

portion of the Chesapeake Bay (1939-1975). Cent. Estuar.

Environm. Sci. Univ. MD 7: 30 p.
Mileikovsky, S. A. 1974. On predation of pelagic larvae and early

juveniles of marine bottom invertebrates by adult benthic

invertebrates and their passing alive through their predators.

Mar. Biol. (Berl.) 26:303-312.

Nelson, T. C. 1957. On the reactions of oyster larvae in relation to
setting on the cape shore of Delaware Bay, N.J. Available from
Dept. Biology, Rutgers Univ., New Brunswick, NJ (unpublished
manuscript)

. 1959. Oyster seed production on Cape May's tidal flats.

Cape May Geographic Soc. Ann. Bull. 13:12-16.

& E. B. Perkins. 1931. Report of the Biology Department.

NJAgric. Exp. Sta. Bull. 522:1-47.
Pritchard, D. W. 1952. Salinity distribution and circulation in the

Chesapeake Bay estuarine system./ Mar. Res. 11:106-123.
. 1953. Distribution of oyster larvae in relation to

hydrographic conditions. Proc. Gulf Caribb. Fish. Inst.

5:123-132.
. 1955. Estuarine circulation patterns. Proc. Am. Soc. Civil

Eng. 81:1-11.
. 1956. A study of the salt balance in a coastal plain

estuary. /. Mar. Res. 15:33-42.
Quayle, D. B. 1969. Pacific oyster culture in British Columbia. Bull.

Fish. Res. Board Can. 169: 34 p.
Seliger, H. H. & J. A. Boggs. 1983. Physical-biological mechanisms

for the transport of oyster larvae in the Chesapeake Bay. Mar.

Biol. (Berl.) 71:57-72.
Sulkin, S. D. (Convenor) 1981. Larval retention in estuaries.

Abstracts for the Sixth Biennial International Estuarine Research

Conference. Estuaries 4:238-240.
Wood, L. & W. J. Hargis, Jr. 1971. Transport of bivalve larvae in a

tidal estuary. Proc. Eur. Mar. Biol. Symp. 4:29-44.

Journal of Shellfish Research, Vol. 3, No. 1, 41-44, 1983.

BIOLOGICAL CONTROL OF FOULING ALGAE
IN OYSTER AQUACULTURE

CATHERINE ENRIGHT, DONNA KRAILO, LARRY STAPLES,
MARIA SMITH, CARL VAUGHAN, DEBRA WARD,
PAMELA GAUL, AND ELISABETH BORGESE

Seafarm Venture, Ketch Harbour
Nova Scotia, Canada BOJ 1X0

ABSTRACT The periwinkle (Littorina littorea Linne) provided excellent biological control of Ectocarpus sp., Entero-
morpha sp., Ulva sp., and pennate diatoms, all of which foul oyster-rearing boxes. The addition of periwinkles (200/m*)
to 1-mm mesh-covered rearing boxes containing juveniles of the European flat oyster Ostrea edulis Linnaes promoted a
significantly higher oyster growth rate (t-test; p = 0.05). Examination of the means obtained from a 5-week study showed
a 30% increase in oyster growth rate when periwinkles were added, in comparison to the unmanipulated control. There was
no significant difference (t-test; p = 0.05) in oyster growth rates when the culture boxes were either brushed once a week
or periwinkles were added. A density range of to 1,600 periwinkles/m of oyster-rearing surface was examined in culture
boxes covered with 6-mm mesh. Similar oyster growth rates were obtained with densities between 300 and 1,600 peri-
winkles/m of oyster-rearing surface. Isopods (Idotea balthica Pallas) at a density of 125/m of oyster-rearing surface were
not effective as a biological control agent.

KEY WORDS: biological control, oysters, periwinkles, algal fouling, Ostrea edulis. Littorina littorea, oyster culture

INTRODUCTION

Oyster-rearing boxes, trays, and lantern nets quickly foul
with algae, mussels, bryozoans, sponges, and other marine
organisms which restrict the flow of water and, consequently,
the availability of phytoplankton to the oysters. Michael
and Chew (1976) examined the effect of progressive fouling
in off -bottom oyster culture in the state of Washington and
correlated it with a decline in the growth rate of the Pacific
oyster Crassostrea gigas Thunberg.

The traditional methods of coping with fouling in oyster
culture include routine manual scraping and brushing,
air-drying, controlled burning, pesticides, and high-pressure
spraying to remove fouling organisms (Arakawa 1980).
Clime and Hamill (1979) found that high -pressure spraying
with a portable 378.5 to 567.7-C/min (100 to 500-gal/min)
capacity pump reduced marine fouling on oyster-culture
gear in Maine. The cleaning schedules included bi-weekly
treatments for small mesh enclosures and monthly cleaning
for lantern nets and larger mesh enclosures during the
height of the growing season. MacLeod (1974) investigated
the use of a hot-water dip treatment for control of fouling
organisms on oyster-culture gear. Huguenin and Huguenin
(1982) examined the use of expanded metal mesh of a
copper-nickel alloy in shellfish trays. Although these proce-
dures are effective, they are both expensive and time
consuming. Dr. E. Scura (Aquatic Farms, Hawaii, pers.
comm.) estimated that 20% of the market price of inten-
sively cultured oysters reflected the costs associated with
reducing fouling organisms during the rearing stages. In
Nova Scotia during 1983, the members of the Ostrea Edulis
Cooperative Association Ltd. allocated more than half of

the labor time associated with rearing oysters to cleaning
of fouling from oysters and culture gear. Thus, fouling has
traditionally been a costly problem in terms of equipment
and labor costs as well as reduced oyster growth rates. An
efficient, inexpensive means of ensuring maximum water
flow about the oysters is greatly needed.

Biological control is the utilization of natural or exotic
species to control the density of undesirable organisms.
Hidu et al. (1981) inadvertently enclosed a rock crab
Cancer irroratus Say in a tray of over-wintering yearling
European oysters and found that the typical thick mat of
fouling organisms did not develop. By selecting crabs of a
distinct size range, Hidu et al. (1981) demonstrated that the
introduction of crabs to oyster culture may provide a means
of biologically controlling the growth of fouling organisms.
Movement by the crab was also believed to reduce silt
accumulation on the oysters. While suitable for the culture
of large oysters, crabs prey upon small oysters and can only
be used with great care as a biological control agent with
juvenile oysters. The fouling problem is more acute with
juvenile oysters because they can not withstand the damage
incurred by traditional cleaning methods. Also, the small-
mesh screen needed to retain juvenile oysters fouls more
quickly and accentuates the fouling problem. Because snails
and isopods have demonstrated the ability to consume algae
(Shaddock and Croft 1981, Steneck and Watling 1982), we
investigated the usefulness of periwinkles and isopods as
biological control agents in juvenile oyster culture. Bequaert
(1943) noted that the herbivorous habits of L. littorea
were sometimes used to keep oysters free of algal growth.
We felt that such an application might be useful in oyster
aquaculture.

ENRIGHT ET AL.

MATERIALS AND METHODS

Juveniles of the European oyster Ostrea edulis Linnaes
were studied inSambro Harbour, Nova Scotia (44°28'5l"N,
63°34'2l"W). The water temperature range was 12 to 17°C
and the salinity range was 29 to 3 1 ppt during the experi-
mental period. The oysters were reared in boxes with
wooden sides which were covered on the top and bottom
with plastic screening. Two sets of three vertically suspended
culture boxes were hung from a floating boom near each
other. The top box in each set was approximately 20 cm
beneath the water surface with subsequent boxes approxi-
mately 25 cm apart. Oyster growth rate was assessed using
change in volume or weight over the experimental period.
An empty box with plastic screen was suspended between
the experimental box sets. A small piece of mesh was
clipped bi-weekly from this box for a microscopic examina-
tion of the colonizing organisms throughout the experi-
mental period. The fouling organisms were identified and
the abundance of each was expressed as a percentage of
the total fresh weight biomass of all fouling organisms.

The first experiment was conducted from 7 July to
12 August 1981. The culture boxes were 83 X 60 X 6 cm
and were covered with 1-mm plastic screening. Each of the
six boxes was divided by wooden slats into four equal
compartments, with each box receiving one of the following
four treatments: the addition of 24 periwinkles (Littorina
littorea) (200/m 2 ) approximately 2 cm in diameter; the
addition of 13 isopods {Idotea balthicd) (125/m 2 ) approxi-
mately 3 cm in length; weekly manual brushing of the
screen mesh; and an unbrushed control. Juvenile oysters,
approximately 5 mm in diameter, were stocked in the
boxes at an initial "density" of 600 g/m 2 .

The second experiment was conducted from 5 July to
3 October 1982. A similarly arranged culture unit was used
with boxes measuring 30 X 30 X 6 cm and covered with

6-mm mesh plastic screen. The six boxes were divided into
four equal compartments and suspended in two units,
each with three boxes. The following series of treatments
was replicated at each of the three-box positions (upper,
middle and lower): weekly manual brushing of the mesh;
(contol), 2, 5, 10, 15, 20 and 25 periwinkles in each
compartment which corresponds to 0.01. 0.03, 0.05,
0.08, 0.10 and 01.3 periwinkJes/m 2 . The oysters used
were approximately 2 cm in diameter and the oyster
stocking "density" was 8,000 g/m 2 .

RESULTS AND DISCUSSION

Littorina littorea proved to be an excellent biological
control agent for reducing algal fouling on the oysters and
on the screens covering the oyster-rearing boxes. The
addition of 200/m 2 periwinkles to 1-mm mesh-covered
rearing boxes containing juvenile European oysters was
shown to yield a significantly higher (t-test; p = 0.05)
oyster growth rate (Table 1). Examination of the means
obtained from a 5-week study showed an approximate
30% increase in oyster growth rate (Set I, 36%; Set II, 25%)
when periwinkles were added compared with the unbrushed
control (Table 1). The major fouling organisms were
Ectocarpus sp. (90%), Enteromorpha sp. (3%), Ulva sp.
(1%), and pennate diatoms (5%). Animal fouling accounted
for less than 1% of the total fouling biomass. There was no
apparent change in the species composition of the fouling
organisms throughout the experimental periods. On the
basis of visual inspections, the periwinkles kept the mesh
cleaner than that obtained with a weekly manual scrubbing.
There was no significant difference (t-test; p = 0.05) in
oyster growth rates when the culture boxes were brushed
once a week or periwinkles were added. Idotea balthica
did not actively graze the fouling organisms which collected
on the plastic screen, and the growth rate of the oysters

TABLE 1.

Increase in volume (m?) and the calculated growth rate (% volume increase day ) of Ostrea edulis cultured in boxes with

unbrushed screens, with brushed screens, with periwinkles, and with isopods. The initial size of the oyster was

approximately 5 mm in diameter and the experimental period was 5 weeks (7 July to 12 August 1981).

Unbrushed

Brushed

With Periwinkles

With Isopod

Box Position

A Volume

% day 1

A Volume

% day '

A Volume

% day '

A Volume

day" 1

Set 1

Upper

190

4.1

240

4.8

240

4.8

170

3.7

Middle

170

3.7

190

4.1

210

4.4

210

4.4

Lower

120

2.7

200

4.2

200

4.2

120

2.7

160

3.5

210

4.4

217

4.5

167

3.6

0.7

0.4

0.3

0.8

Set II

Upper

260

5.0

280

5.2

310

5.5

260

5.0

Middle

220

4.5

320

5.6

300

5.4

200

4.2

Lower

180

3.9

190

4.7

220

5.2

140

3.8

220

4.5

263

5.2

277

5.4

200

4.3

0.6

0.5

0.2

0.6

Biological Control of Fouling algae

reared in such compartments did not differ significantly
(t-test; p = 0.05) from that of the oysters in the unbrushed
(control) compartments. Using a comparable isopod density.
Shaddock and Doyle (1983) found that /. balthica vora-
ciously grazed Ectocarpus sp., a brown seaweed which
grows epiphytically on Chondrus crispus in tank cultures.
Perhaps in the present experiment a higher isopod density
would have negated the fouling rate in the oyster-rearing
boxes. Oyster boxes suspended in the water column may
not provide an adequate habitat for isopods; perhaps their
feeding behavior is altered in that setting. From the data
in Table 1, it is clear that higher oyster growth rates were
obtained in box Set I compared to box Set II. The difference
may have been the result of their relative position in the
bay as box Set II was downstream from box Set I with
respect to the food source. All other parameters were the
same in each box set.

An examination of a periwinkle density range from to
1 ,600/m 2 of mesh-rearing surface, when a 6-mm mesh size
was used, indicated little change in oyster growth rates

between 300 and 1 ,600 periwinkles/m 2 of screen (Figure 1 ).
The optimal periwinkle density would be expected to vary
as a function of the degree of fouling and with factors that
influence the periwinkle grazing rate (e.g., temperature).

There are many advantages to utilizing periwinkles for
biological control of fouling organisms in juvenile oyster
culture. Periwinkles are herbivors; therefore, they do not
prey on oysters as do crabs and other organisms. Littorina
littorea is extremely abundant in western Europe and in
northeastern North America and locally exceed densities
of 150 periwinkles/m 2 in the low intertidal zone. The
periwinkle can completely withdraw its soft tissue into its
shell, thus protecting itself against desiccation when the
oyster boxes are removed from the water for data collec-
tion or transportation. There was no evidence of erosion
of the mesh fibers as a result of the periwinkles grazing
along the plastic screens. The major advantage of using a
biological control agent such as a periwinkle is the reduction
in costs associated with cleaning algal fouling organisms. As
water flow and phytoplankton availability are greatly

500-

400-

300-

5 200

<u
to
o
oj

o 100

-5.0

400

200

800

Periwinkles • m
on oyster mesh rearing surface

-2

-rV/ 1

1600 BRUSHED
WEEKLY

4.0

3.0

•*—

2.0

-♦—

*—

1.0

CJ5

Figure 1. Increase in weight (g fresh weight) and the corresponding calculated growth rate {% weight increase day" 1 ) of Ostrea edulis
cultured with Littorina littorea at various densities and compared with a weekly, manual mesh-brushing treatment. The initial size of the
oysters was approximately 2 cm in diameter and the mesh used on the rearing boxes was 6 mm. The experimental duration was 12 weeks
(5 July to 3 October 1982). Standard deviations are shown (n = 3).

ENRIGHTETAL.

enhanced for juvenile oysters cultured with periwinkles,
the need to transfer oysters on to larger mesh sizes, as is
presently the practice (Clime and Hamill 1979), is reduced.
Such cost reductions will greatly improve the profitability
of off-bottom oyster culture.

ACKNOWLEDGM ENTS

The financial assistance from the Nova Scotia

Department of Development, Provincial Employment Pro-
gram, is gratefully acknowledged. We thank P. Shacklock,
S. Smith and J. Dale for their assistance on site. Sincere
appreciation is expressed to Drs. J. Craigie, G. Newkirk,
and H. Hidu for reviewing the manuscript. This study is
dedicated to the memory of T. Moore, who assisted greatly
in the initial stages of this project.

REFERENCES CITED

Arakawa, K. Y. 1980. Prevention and Removal of Fouling on
Cultured Oysters: A Handbook for Growers. Translated from
Japanese by R. Gillmore. Univ. Maine Sea Grant Tech. Rep.
No. 56: 56 p.

Bequaert, J. C. 1943. The genus Littorina in the western Atlantic.
Johnsonia 7:1-28.

Clime, R. & D. Hamill. 1979. Growing oysters and mussels in Maine.
Golden, E., ed. Aquaculture Development Workshop; Bath, ME:
Coastal Enterprises. Inc. 46 p.

Hidu, H., C. Conary & S. R. Chapman. 1981. Suspended culture of
oysters: biological fouling control. Aquaculture 22:189-192.

Huguenin, J. E. & S. S. Huguenin. 1982. Biofouling resistant shell-
fish trays. J. Shellfish Res. 2(l):41-46.

Michael, P. C. & K. K. Chew. 1976. Growth of Pacific oysters.

Crassostrea gigas, and related fouling problems under tray

culture at Seabeck Bay, Washington. Proc. Natl. Shellfish. Assoc.

66:34-41.
MacLeod, L. L. 1974. Controlling blue mussel (Mytilus edulis)

fouling on oysters and oyster trays with hot water immersion.

8 p. Unpublished document. Available from: Nova Scotia Dep.

fish.. Resour. Develop. Div. Fish. Train. Cen. Pictou, NS, Canada.
Shacklock, P. F. & R. W. Doyle. 1983. Control of epiphytes in

seaweed culture using grazers. Aquaculture 31:141-151.
Shacklock, P. F. & G. C. Croft. 1981. Effect of grazers on Chondrus

crispus in culture. Aquaculture 22:331-342.
Steneck, R. S. & L. Watling. 1982. Feeding capabilities and limita-
tion of herbivorous molluscs: a functional group approach.

Mar. Biol. (Berl.j 68:299-319.

Journal of Shellfish Research, Vol. 3. No. 1, 45-50, 1983.

A STUDY OF GLUCOSE, LOWRY -POSITIVE SUBSTANCES, AND

TRIACYLGLYCEROL LEVELS IN THE HEMOLYMPH OF

CRASSOSTREA VIRGINICA (GMELIN)

MARY L. SWIFT AND MOHAMMED AHMED

Department of Biochemistry
College of Medicine
Howard University
Washington, DC 20059

ABSTRACT Oysters, Crassosrrea virginica (Gmelin), were maintained in the laboratory under controlled conditions
of temperature and salinity. Levels of several hemolymph constituents were analyzed. Average values of hemolymph glucose,
Lowry-positive substances, and triacylglycerols were 8.83 ± 1.98 mg/100 mC (± SE), 11.0 ± 1.89 rag/mf (± SE), and
43.2 /Jg/100 mC, respectively. Hemolymph glucose values varied over a wide range. No deleterious effects of this variance
(as judged by mortality rates) could be detected. Groups of animals with initial hemolymph glucose levels of 23.1 to
25.0 mg/100 m? survived as long as those with initial values of 5.3 to 8.4 mg/100 mC. Oysters held at constant water
temperatures and salinities tended to maintain the concentration of their hemolymph glucose and Lowry-positive substances
over a 27-day period of starvation; hence, some type of regulatory mechanism is involved in controlling the levels of these
metabolites in oyster hemolymph. Extremes in environmental conditions appear to affect the concentrations of these
metabolites in hemolymph. Groups of oysters maintained in sea water at a temperature of 4 C had significantly higher
(p < 0.05) levels of hemolymph glucose and Lowry-positive substances than groups held at 20 C. Groups of oysters
maintained at alow ambient salinity (12 ppt) had significantly lower (p ^0.05) levels of hemolymph glucose and Lowry-
positive substances than groups kept in water of 18 ppt and 24 ppt salinity.

KEY WORDS: oyster, Crassostrea virginica, hemolymph, glucose, regulation

INTRODUCTION

Traditionally, the physiological and nutritional condi-
tions of oysters have been monitored by evaluating tissue
glycogen content (Gabbott and Walker 1971. Willis et al.
1976). The deposition and utilization of not only glycogen
but also lipid by the American oyster may be influenced by
a number of factors. Seasonal variations in tissue glycogen
and lipid content, which are keyed to the reproductive
cycle, are well documented (Galtsoff 1964, Krishnamoorthy
et al. 1979, Swift et al. 1980). The effects of starvation on
these metabolic reserves in oysters have been examined
(Riley 1976, Willis et al. 1976, Swift et al. 1980), as have
environmental conditions which may also affect the rate of
synthesis or utilization and, therefore, content of metabolic
reserves.

Several groups have investigated either the whole
animal response or the response of selected excised tissues
to changes in temperature and salinity. Ruddy et al. (1975)
examined the growth rate of Crassostrea virginica (Gmelin)
during exposure to a warm water temperature ( 14 to 19°C).
Levels of each of the major classes of metabolites (carbo-
hydrate, protein, and lipid) increased in these animals. At
the same time gonadal development occurred four months
earlier than usual. Similar increases in biochemical reserves
have been observed in Crassostrea gigas (Thurnberg) and
Ostrea edulis (Linne) (Mann 1979). Percy and Aldrich
(1971), Percy et al. (1971), and Bass ( 1977) monitored the
effect of changes in ambient water temperature and salinity

on oxygen consumption of excised gills, mantle, and
adductor muscle of C. virginica. These reports agree that,
with increasing temperature or decreasing salinity, oxygen
use increases. When subjected to extremes of temperature
and salinity, these animals used more oxygen (Shumway
and Koehn 1981). These data imply that the metabolic
rate has increased and, thus, utilization of metabolic
reserves has increased, resulting in a decrease in tissue
content of glycogen and lipid.

Despite the proven usefulness of data on tissue composi-
tion, the processes required to obtain them are cumbersome
and time consuming. In contrast, more complete information
concerning the nutritional and physiological conditions of
mammalian organisms may be obtained easily and rapidly
by analysis of blood metabolites. Unfortunately little is
known regarding the metabolite levels in the hemolymph of
C. virginica. Hand and Stickle (1977) studied the effect of
tidal-like fluctuations in salinity of ambient sea water on
pericardial fluid composition of the oyster. Ion concentra-
tions, except K\ were found to be isoionic to the various
ambient salinity regimes: ninhydrin-positive substances
ranged from 1 .5 to 6.0 mM.

The lack of suitable data in the literature for establishing
baseline values for hemolymph glucose, protein, and triacyl-
glycerol levels in C. virginica prompted the following studies.
Glucose," total Lowry-positive substances (LPS), and triacyl-
glycerols were examined in hemolymph from groups of
oysters subjected to: (1) starvation, (2) different ambient
temperatures, and (3) different ambient salinities.

SWIFT AND AHMED

MATERIALS AND METHODS

Oysters (C. virginica), purchased commercially (Capt.
White and Sons, Seafood, 110 Main Avenue, SW, Washing-
ton, DC 20024), had been harvested two or three days
before arrival in the laboratory. The height of the animals,
measured as the distance from the hinge to the extreme
ventral margin of the shell, ranged from 7 to 1 2 cm. Before
any data were gathered the oysters were cleansed in tap
water with the aid of a wire brush and acclimated to
laboratory conditions for three days. Up to 20 unfed
individuals were held in an aquarium in approximately 7 C
of artificial sea water (Instant Ocean, Aquarium Systems
Inc., 33208 Lakeland Blvd., Eastlake, OH 44094). The
glass holding tanks were arranged so that the sea water was
drawn off at the bottom of each tank, and then pushed up
through a water-cooled condenser to the top of the holding
tank by compressed air (Swift et al. 1975). A refrigerated
bath and circulator was used to control the water tempera-
ture. Sea water in the tank was changed every two days
and the tank thoroughly rinsed at those times.

Hemolymph was collected with a small syringe from die
pericardial cavity of carefully opened oysters. The hemo-
lymph was placed in an ice-cooled centrifuge tube. Cellular
debris were separated from the hemolymph by centrifuga-
tion at 1,000 X g for 20 minutes at 4°C. The supernatant
liquid was transferred to a small vial and stored at — 10°C
before glucose, total Lowry-positive substances (LPS), and
triacylglycerol determinations were accomplished. Glucose
was analyzed using the glucose oxidase method (Bergmeyer
and Bernt 1974), total Lowry-positive substances were
estimated according to Lowry (Lowry et al. 1951), and
triacylglycerol was analyzed by the acetylacetone test
(Fletcher 1968) with a slight modification. Hemolymph
that was pooled from 3 to 4 oysters was extracted with
n-heptane; 1 m2 of the upper layer was removed for analysis.
After the aliquot was dried completely under a stream of
air, 2.0 m2 of isopropanol were added. Thereafter the
procedure was the same as described by Fletcher (1968).

Hemolymph lipids were extracted by the Folch proce-
dure (Folch et al. 1957). The chloroform layer, remaining
after the aqueous NaCl wash, was evaporated to dryness
under reduced pressure. The lipids were redissolved in a
minimal quantity of 2:1 (v/v) chloroform :methanol and
separated by thin-layer chromatography on silicic acid
using n-hexane:diethyl ethenglacial acetic acid at a volu-
metric ratio of 70:30:1 (Malins and Mangold 1960). The
spots were visualized by iodine vapor retention or by
ultraviolet fluorescence after spraying the chromatogram
with 0.2% V :7'-dichlorofluorescein in 95% ethanol.

To examine the effect of selected environmental condi-
tions on the levels of metabolites in oyster hemolymph,
groups of unfed animals were held in tanks for up to
27 days under the following conditions: (1) in 24 ppt sea
water at temperatures of 4, 10, 15, or 20°C, and (2) in 12,

18, or 24 ppt sea water at 20° C or 15°C. Data were analyzed
for significance (p < 0.05) by the Student's /-test.

RESULTS

Oysters obtained throughout the course of this study did
not have significantly different initial levels of hemolymph
glucose (Table 1). Overall hemolymph glucose concentra-
tions averaged 8.83 ± 1.98 mg/100 mC (± SE) and ranged
from 1.9 to 25.0 mg/100 ml. Hemolymph LPS levels
averaged 11.0 ± 1.89 mg/m2 and ranged from 3.17 to
29.5 mg/mE. Hemolymph triacylglycerol values were quite
low averaging 43.2 /ug/100 mC and ranged from 3.3 to
200 Mg/100 m8.

TABLE 1.

Initial hemolymph glucose, Lowry-positive substances (LPS)
and triacylglycerol levels in groups of oysters.

Glucose

LPS

Triacylglycerol

Month

(mg/100 mC)*

(mg/mS)*

(Ag/lOOmC)**

December

15.80 ±6.54

26.00 ±4.18

11.7

January

9.18 ±2.18

18.60 ±3.18

25.0

February

8.96 ±1.46

19.40 ±4.18

15.6

March

12.90 ±2.28

14.10 ± 1.30

—

April

108

8.41 ±2.50

12.10 ±2.34

26.3

May

3.14 ±1.22

8.08 ±1.41

30.0

June

9.09 ±2.11

8.02 ±2.37

43. 9f

*Mean values ± SE
**Mean values obtained by pooling hemoymph from 3 or more
individuals
t76.7 jug/100 m£ if values of 150 and 200 jUg/100 m£ are included

No free or nonesterified fatty acids could be detected
in oyster hemolymph using standard analytical techniques
or after lipid extraction followed by thin-layer chromatog-
raphy. This is in agreement with results of other lipid
analyses of oyster tissues (Watanabe and Ackman 1977,
Bunde and Fried 1978, Ghassemieh 1978).

Oysters held at constant temperature and in sea water
of constant salinity tended to maintain their hemolymph
glucose, LPS, and triacylglycerol concentrations over a
27-day period of starvation (Tables 2, 3. and 4); however,
extremes in external conditions appear to affect the concen-
trations of these metabolites. Groups of unfed oysters
maintained in 24 ppt artificial sea water at temperatures of
4°C had significantly higher (p < 0.05) levels of hemolymph
glucose and LPS when compared to values obtained from
oysters kept at 20°C. Oysters held at 4°C had hemolymph
glucose values of 19.3 ±3.5 mg/100 mC while those kept at
20°C had hemolymph glucose values of 8.41 ± 1.4 mg/
100 mC. Similarly the mean LPS values were 17.56 ±
1.42 mg/mC and 9.76 ± 0.85 mg/mC for the animals at
4°C and 20°C, respectively. At a low ambient salinity of
1 2 ppt, oyster hemolymph glucose and LPS concentrations
were significantly (p < 0.05) decreased when compared to
the values found in oysters kept in water of 18 and 24 ppt
(Tables 5 and 6).

Study of Oyster Hemolymph

TABLE 2.
Hemolymph glucose levels* (mg/100 m?) in starved oysters maintained in 24 ppt sea water at different temperatures.

Temperature ( C)

Number of Days

23.4 ±11.2

23.1 ± 10.6

13.2 ± 2.59a
19.6 ± 4.07(5)

13.3 ±10.0(3)3

19.3 ± 3.523

9.18 ±2.17
10.7 ±2.04(5)
11.3 ±1.74

7.72 ±1.83

6.63 ±1.75

11.7 ±3.81

10.2 ±2.81

5.33 ± 1.19

8.38 ±2.14
6.43 ±1.25 b
13.3 ±6.02(5)
5.92 ± 0.904b

Group Mean

10.3 ±l.ll b

9.06 ± 1.33b

8.44±1.43 b

*Mean value obtained from six individuals ± SE, unless otherwise indicated. Number in parenthesis shows number of oysters used. Means
assigned the same or no superscript were not significantly different. Means assigned different superscripts were different at p ^0.05 level
(compared across groups).

TABLE 3.
Hemolymph Lowry -positive substance levels* (mg/m?) in starved oysters maintained in 24 ppt sea water at different temperatures.

Temperature ( C)

Number of Days

7
14
24

26.0 ±4.18 a

15.1 ±1.52
14.1 ± 1.17
17.1 ±1.17(5)3
12.3 ±1.64(3)

17.5 ± 1.42 a

18.6 ± 3.18

14.5 ±2.86
15.1 ±1.68

19.4 ±4.18 (5)
16.8 ±1.32(5)

21.0 ±4.59
17.7 ±4.59

10.1 ±2.23(3)b
12.1 ±2.02(5)

6.7 ±1.23
11.1 ±1.84(5)b

8.71 ±1.93

Group Mean

16.1 ±2.57

18.7 ±3.56

9.76 ±0.85 b

'Mean value obtained from six individuals ± SE, unless otherwise indicated. Numbers in parenthesis show number of oysters used. Means
assigned the same or no superscript were not significantly different. Means assigned different superscripts were different at p ^0.05 level
(compared across groups).

TABLE 4.

Hemolymph triacylglycerol levels* (JLlg/100m6) in starved oysters
maintained in 24 ppt sea water at different temperatures.

Temperature ( C)

Number of Days

11.7

25.0

6.25

16.9

13.4

23.8

6.25

55.0

8.33

6.25

47.5

6.25

—

25.0

113.0

—

27.3

*Pooled samples from 3 to 6 oysters.

DISCUSSION

Hemolymph glucose levels have been examined in other
fasting molluscan species. In the terrestrial snail, Stropho-
cheilus oblongiis (Miiller), hemolymph glucose values
ranged from 2.5 mg/100 mx 1 to 16.88 mg/100 m2 (Marques
and Falkmer 1976). Hemolymph glucose levels in the
freshwater pulmonate snail, Lymnaea stagnalis jugidaris
(Say), ranged from 1.86 to 5.68 mg/100 m2 (X = 3.0) and

1.9 to 4.0 mg/100 mx 1 (X = 2.9) in separate investigations
(Friedl 1968, 1971 ). Hemolymph glucose concentrations in
two freshwater bivalve molluscs, Anodonta cygnea (Linne)
and Unio pictorum (Linne) averaged 9.4 ± 0.49 mg/100 m2
and 14.0 ± 1.6 mg/100 mS, respectively (Plisetskaya et al.
1978). The hemolymph glucose level in the Atlantic deep
sea scallop, Placopecten magellanicus (Gmelin). was 2.6 ±
0.6 mg/100 mC (Thompson 1977); and the hemolymph
glucose concentration in another marine bivalve, Mytilus
edulis Linne, lies between 16.0 and 37.0 mg/100 m?
(Bayne 1973).

Inspection of these data leads to the conclusion that
hemolymph glucose values during fasts in several molluscan
species may vary over a wide range and are not directly
related to terrestrial, freshwater or marine habitats. Thus,
it may be inferred that these animals, including the oyster
C. virginica, are more tolerant of larger variations of glucose
concentrations in circulatory fluids than mammals. In this
study, no deleterious effects of variations in hemolymph
glucose levels could be detected. Groups of oysters with
initial hemolymph glucose levels of 23.0 to 25.0 mg/100 mx 1
survived as long as those with initial hemolymph glucose
values of 5.3 to 8.5 mg/100 mE.

SWIFT AND AHMED

TABLES.

The effect of ambient water salinity on hemolymph glucose levels* (mg/100 m£) of starved oysters.

Temperature ( C)

Salinity (ppt)

Number of Days

3.14 ±1.22(5)
2.46 ±0.47(5)
3.98 ±0.69

7.08 ±2.58(5)
3.74 ±0.92(5)
2.88 ±0.75(5)
3.72 ±0.93(5)

5.33 ±1.19
8.38 ±2.14
6.43 ±1.25
13.3 ±6.02(5)
5.92 ±0.90

6.72±0.92 b

10.2 ±1.09(5)
7.26 ±0.88(5)
2.16 ±0.51(5)
4.58 ±0.97(5)

7.87 ±1.82

6.63 ±1.76

11.7 ±3.82

10.2 ±2.86

Group Mean

3.24 ±0.59

4.57 ±1.00

6.53±1.00 b

9.02 ± 1.58 b

*Mean values obtained from six individuals ± SE. Number in parenthesis shows number of oysters used. Means assigned the same or no
superscript were not significantly different. Means assigned different superscripts were different at p ^0.05 level (compared across groups).

TABLE 6.

The effect of ambient water salinity on hemolymph Lowry-positive substance levels* (mg/m?) of starved oysters.

Temperature ( C)

Salinity (ppt)

Number of Days

7
14
24

8.08 ±1.41
5.98 ± 1.70(4) a
8.68±0.96(4) a

12.2 ±2.41(3)
6.51 ±0.73(4) a
9.99 ±1.99(4)

10.1 ±2.23(5)
12.1 ±1.23(5) b
6.73 ±1.23
11.1 ±1.84(5)

3.17 ±0.71(5) a
9.48 ± 1.74
7.68 ±1.94(5)

19.4 ±4.18(5)
16.8 ± 1.32 b

18.3 ±4.61

17.7 ±4.15

Group Mean

6.07 ±0.91 a

9.31 ± 1.15 b

9.64 ± 1.17 b

6.78 ± 1.15 b

17.5 ±2.10 b

During the course of these studies hemolymph glucose
levels were relatively stable within test groups. This indicates
that some type of regulatory mechanism functions in the
oyster. There is no direct evidence for the regulation of
hemolymph glucose in other molluscs; however, indirect
evidence concerning various aspects of this physiological
mechanism has been published. Enzymatic activities which
are necessary for the postulated regulation have been
identified in several molluscs. For example, hexokinase and
glycogen phosphorylase activities have been reported in
Pecten maximus (Linne), O. eclulis, Ensis ensis flinne),
Chlamys varius (Linne) (Zammit and Newsholme 1976),
and C. gigas (Nakamuro et al. 1980). Glycogen synthase
activity has been studied in M. edulis (Cook and Gabbott
1978, Gabbottet al. 1979).

Of the hormones known to affect mammalian blood glu-
cose levels, only insulinhasbeen investigatedin some molluscs.
Hemolymph glucose levels in A. cygnea, U. pictorum
(Plisetskaya et al. 1978), and S. oblongits (Marques and

Falkmer 1976) are affected by insulin in ways analogous to
those found in mammals. In addition, insulin-like proteins
have been reported in several freshwater bivalves (Pliset-
skaya et al. 1978), a terrestrial snail (Marques and Falkmer
1976). and in saltwater bivalves (Collip 1923, Fritsch
and Sprang 1977), including O. edulis (DeMartinez et al.
1973).

Hemolymph triacylglycerol levels in two other bivalves
were at least 20 times those found in oysters in this study.
Triacylglycerol concentration in the hemolymph of the
hard clam, Mercenaha mercenaria (Linne), was 1 mg/100 m?
(Hoskin and Hoskin 1977), and in the plasma of the deep-
sea scallop,/ 1 , magellanicus, values ranged from 0.1 to 1 mg/
100 m5 (Thompson 1977). The low levels of hemolymph
triacylglycerols and free fatty acids in bivalve molluscs may
be a consequence of their general metabolic strategy. As
facultative anaerobes (Zandee et al. 1980) these animals
would be more dependent upon carbohydrate for energy
than lipid.

Study of oyster Hemolymph

Few reports on the concentration of hemolymph proteins
have appeared. Hand and Stickle (1977) examined ninhydrin-
positive substances in whole hemolymph from C. virginica.
Their values ranged from 193 to 702 mg/mC; however,
those investigators were studying hemolymph which had
not been subjected to centrifugation and. in addition, the
ninhydrin method detects not only protein but also free
amino acids. Thus, the large differences in data from the
two laboratories may be explained. On the other hand,
plasma from P. magellanicus contained LPS in the range of
1.55 to 2.17 mg/mB (Thompson 1977).

The different levels of hemolymph glucose and LPS
which were observed after the oysters were exposed to
several temperature and salinity regimes may reflect adaptive
metabolic mechanisms. These adaptive mechanisms would
be necessary because oysters are sessile and, thus, subjected
to the challenges of a changing euryhaline habitat. For
example, successful acclimation to changing ambient
salinity is apparently closely related to hemolymph amine
concentration. Other investigators have found that hemo-
lymph protein and amino acid levels not only in C. virginica
(Hand and Stickle 1977), but also in Pyrazus ebeninus
(BruguiJre) (Ivanovici et al. 1981) as well as the tissue free
amino acid values (Lynch and Wood 1966), vary directly

with ambient salinity. This phenomenon was readily
observed with ambient salinity changes of > 6 ppt provided
that the animals had been acclimated to the particular
salinity for a period of at least two weeks. This is the first
report that hemolymph glucose levels also vary with ambient
salinity.

Temperature also affects the metabolism of bivalve
molluscs. Oysters that are held at elevated temperatures
have increased metabolic rates as measured by increased
oxygen utilization (Percy and Aldrich 1971, Percy et al.
1971, Shumway and Koehn 1981). As ambient tempera-
ture increases, oyster hemolymph glucose levels decrease.
Similarly short-term exposure (30 to 60 hours) of My tilus
galloprovinciallus Lamarck to elevated temperature regimes
caused a decrease in hemolymph glucose (Madar et al.
1980). The physiological importance of these findings
remains to be explored.

ACKNOWLEDGMENTS

This work was supported in part by a Biomedical
Research Support Grant No. 5S07. RR03561. from the
General Research Support Branch, Division of Research
Resources, National Institutes of Health. Bethesda, MD.

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Journal of Shellfish Research, Vol. 3, No. 1, 51-57. 1983.

EFFECT OF RATION ON GROWTH AND GROWTH EFFICIENCY
OF JUVENILES OF CRASSOSTREA VIRGINICA (GMELIN)

EDWARD R. URBAN, JR., GARY D. PRUDER
AND CHRISTOPHER J. LANGDON

Center for Mariculture Research
University of Delaware
Lewes, Delaware 19958

ABSTRACT Juveniles of Crassostrea virginica were batch-fed on different rations of an algal diet of Tlialassiosira
pseudonana and Isochrysis aff. galbana in experiments lasting three weeks and the resulting growth and growth efficiencies
were determined. Maximum growth occurred when the oysters were fed on the highest daily ration tested which was equal,
at the beginning of an experiment, to an algal dry weight of 4.6% of oyster live weight. Weight-specific rations decreased
during each week of growth experiments because rations were only adjusted for oyster growth on a weekly basis. An
initial daily ration of 4.6% was calculated to be equivalent to an effective daily ration of 2.8% of oyster live weight or
59.6% of oyster dry organic weight per week of an experiment. Highest growth efficiencies of 17.9 to 22.6% occurred with
effective rations of 1.4 to 2.3% of oyster live weight. The experimental results indicated that weekly adjusted rations
based upon previously reported formulae for the prediction of adequate rations for C. virginica may not be sufficient in
meeting the requirements of juvenile oysters for maximum growth.

KEY WORDS: ration, oyster, growth, algae, growth efficiency, Crassostrea virginica

INTRODUCTION

Successful rearing of bivalve molluscs for both research
and commercial purposes depends upon the delivery of an
adequate food ration. Despite many attempts to develop
satisfactory nonalgal diets or supplements (e.g., Chanley
and Normandin 1967, Winter 1974, Masson 1977, Epifanio
1979), algae remain indispensable as the principle food
source for artificially reared bivalves. Growth studies have
resulted in the determination of the relative food qualities
of different algal species (for reviews see Epifanio [1983]
and Webb and Chu [1983]); however, the relationship
between ration size and bivalve growth rate has not been
adequately studied for many bivalve species.

The most complete studies on the relationship between
ration size and growth of bivalves were conducted by Bayne
and co-workers with Mytilus edulis L. (Bayne 1976,
Widdows 1978a,b), and Navarro and Winter (1982) for
Mytilus chilensis Hube. On the basis of measurements of
the energy balance of Mytilus spp. fed on a range of algal
rations under different conditions of algal cell density and
animal body weight, numerical relationships were formu-
lated that integrated these variables in a predictive model of
"scope for growth." Scope for growth can be defined as the
energy of the assimilated ration available for somatic and/or
germinal tissue growth, once metabolic energy requirements
have been met (Warren and Davies 1967). Bayne and Worall
(1980) and Navarro and Winter (1982) found close agree-
ment between growth of mussel populations in the field
and growth predicted by such mathematical models. Less
is known about the interrelationships among ration, metabo-
lism, and growth for oysters, although assimilation and
growth efficiences of Crassostrea virginica (Gmelin) have
been reported by several workers (Tenore and Dunstan

1973, Langfoss and Maurer 1975, Romberger and Epifanio
1981, Valenti and Epifanio 1981).

Predicting optimum algal rations for maximum oyster
growth on the basis of caloric measurements and scope for
growth determinations is of limited practical usefulness
because algal diets vary in their nutritive value (Epifanio
1983, Webb and Chu 1983); thus, an algal ration may be
calorifically satisfactory but biochemically deficient in
some essential nutrient for growth. Because factors deter-
mining algal food value are not fully understood, optimum
rations for maximum oyster growth must be determined
empirically.

In this study, the effect of algal ration on the growth
and gross growth efficiency of juveniles of C. virginica was
determined. The tested rations were compared with the
predicted rations for maximum oyster growth described by
Epifanio and Ewart (1977), Pruder et al. (1977), and
Epifanio (1979).

MATERIALS AND METHODS

Juveniles of C. virginica were fed different algal rations
in a series of four experiments. In each experiment, groups
of 20 oysters were randomly chosen from a population of
similar sized oysters. Initial oyster live weight did not vary
by more than one standard deviation of the population
mean live weight. The identities of individual oysters were
maintained during growth experiments by partitioning the
oysters in 400 fim mesh trays, which were submerged in 4 2
of l-/im-filtered seawater at 30 ppt salinity and 25 C. The
cultures were aerated to keep the algal cells in suspension
and the seawater was changed daily.

The animals were fed rations composed of a 50/50 mix-
ture (based on dry weight [wt] )of Thalassiosira pseudonana

URBAN ET AL.

Hasle and Heimdal (clone 3H) and Isochrysis aff. galbana
Parke (clone T-ISO). This algal mixture supports excellent
growth of juveniles of C. virginica (Ewart and Epifanio
1981). The algae were cultured in 250-8 containers at 19°C,
illuminated with 550-600 /iW/cm 2 of light (cool white
fluorescent lamps), and nutrient enriched with f/2 medium
(Guillard 1975). Algal cell dry weights were assumed to be
1.32 X 10~ 8 mg/cell for T. pseudonana (Epifanio and Ewart
1977) and 2.01 X 10" 8 mg/cell for/, aff. galbana (S. Ali, Uni-
versity of Khartoum, Port Sudan, Sudan, pers. comm.). Algal
concentrations were determined using a hemocytometer.

Initial algal rations that ranged in dry algal weight from
0.52 to 4.6% of oyster live weight were tested in growth
experiments (Table 1). Algal concentrations ranged from
0.12 mg dry wt algae/C (10,000 cells/mC) to 2.60 mg dry
wt algae/6 (217,000 cells/mx 1 ) (Table 1). By adding one-half
the algal ration twice a day to the 4-2 culture vessels, it was
possible to feed oysters algal cell concentrations which
never exceeded 500,000 cells mC" 1 , and, therefore, were less
than concentrations reported to cause pseudofecal produc-
tion in C. virginica (Epifanio and Ewart 1977). Clearance
of algal cells was greater than 95% per day in all treatments
and, therefore, little loss of ration occurred.

Oysters were weighed individually at the beginning of
each experiment. Group live weights were used for weekly
adjustments of rations to compensate for oyster growth
during each week of the experiment. At the end of the

experiments, oysters were reweighed individually, dried to
constant weight at 60°C, weighed, and then ashed at 450°C
for 24 to 48 hours and reweighed (Walne and Millican 1978).
The difference between total dry weight and ash weight was
assumed to be equal to total oyster organic weight. Individual
live, dry, ash, and organic weights were similarly determined
for an initial sample of 50 oysters at the beginning of each
experiment.

RESULTS

The weight-specific daily rations decreased during each
week of an experiment as a result of the growth of the
animals and because the rations were only adjusted weekly
(Figure 1). This decrease was greatest in treatments with
rapidly growing oysters. To obtain a better estimate of the
effective ration fed to the oysters, the geometric mean of
the actual daily ration was determined for each week of an
experiment. The overall effective ration for the 3-week
experiment was calculated as the mean weekly effective
ration (Table 1 ).

Oyster growth rate increased with increasing effective
ration over the range tested of 0.2 to 2.8% of oyster live
weight (Table 1 and Figure 2). The highest effective algal
ration of 2.8% of oyster live weight was equivalent to a
ration of 59.6% of oyster dry organic weight, based on a
mean dry organic contentof 4.7%foroystersfromtwoexperi-
ments (Table 2). Regression analysis of log-transformed.

TABLE 1.

Initial, final, and effective percent rations and the resulting growth
of juveniles of Crassostrea virginica after 3 weeks.

Initial Ration Concentration
(mg dry wt algae Z 1 )

Percent Rations*

Initial

Final

Effective

k Value

2.60

4.6

1.9

2.8

0.128

2.60

4.6

1.9

2.8

0.123

1.95

3.5

1.6

2.3

0.107

2.60

3.3

1.7

2.2

0.098

1.30

2.3

1.2

1.6

0.093

1.30

2.3

1.2

1.6

0.091

0.97

1.7

1.0

1.2

0.070

2.60

1.9

1.2

1.4

0.067

0.65

1.2

0.8

0.9

0.057

1.95

1.4

1.0

1.1

0.053

0.65

1.2

0.8

1.0

0.049

0.65

0.8

0.6

0.037

1.30

0.9

0.7

0.037

0.32

0.6

0.5

0.027

0.65

0.5

0.4

0.018

0.12

0.2

0.013

unfed

0.0

0.009

unfed

0.0

0.005

unfed

0.0

0.003

Percent Increase in Oyster Live Wt

1363

1226

847

687

604

585

338

305

231

203

183

120

107

22t

10*

*Percent ration = ([dry wt of algae per oyster live wt] X 100). Effective ration is the geometric mean ration for each week, averaged for the

3-week experiment (Figure 1).
fk is the daily instantaneous relative growth rate (see RESULTS for formula).
jLive weight increases of unfed oysters probably resulted from increases in inorganic shell weight because the organic content of unfed

oysters decreased during the experiment (Table 2).

Effect of Ration on Growth of Juvenile Oysters

weekly live-oyster weights plotted against time, indicated
that growth occurred at a constant exponential rate for
oysters fed on the 2.8% effective ration (r 2 = 0.997, F,
2615.3, p < 0.001).

(1.6)

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1600
1200
800 H
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F = 26I5.3 p<0.00l
(1,6)

the effective % ration

(geometric mean of actual % ration)

7 14 21

ELAPSED TIME -DAYS

Figure 1. Change in percent daily ration for Crassostrea virginica fed
an initial daily ration of 4.6%. The upper curve shows the growth of
juveniles of C. virginica fed an initial daily ration of 4.6% over a 3-
week period. The lower figure shows the change in the percent daily
ration over the course of each week of the experiment. The vertical
arrow indicates the effective ration for each week. The t-value in
the exponential equation is in days.

The daily instantaneous relative growth rate (k) was
calculated for each ration (Table 1 ). where

k= [(dWt/dt)/Wo] = (2.303/t) log (Wt/Wo)

and Wo is the initial live weight (mg) and Wt is the final
live weight (mg) after 21 days (t) of growth (Brody 1945).
The k values and values for percentage increase in oyster
live weight were both directly dependent on the weight-
specific ration and were not greatly affected by the concen-
tration of algae added to obtain the required ration (Table 1).

— 14

O
><

c
«
o

Effective
% Ration

1 2 3

elapsed time (weeks)

Figure 2. Increase in live weight of juveniles of Crassostrea virginica
fed different effective percent rations of an algal diet of Thalassiosira
pseudonana and Isochrysis aff. galbana. Percent increase in live
weight was calculated from initial live weight.

Table 2 and Figure 3 give gross growth efficiencies for
oysters fed different rations. Figure 3 shows that gross
growth efficiency increased from -37.7% at an effective
ration of 0.2% to a maximum of 22.6% with an effective
ration of 1.4%. Gross growth efficiency declined slightly
as rations were increased from 1.6 to 2.8%. From Figure 3,
the maintenance ration for juvenile oysters cultured under
the described conditions was 0.5% of oyster live weight.
The organic content of both starved oysters and oysters fed
a 0.2% effective ration decreased over the experimental
period, compared with initial samples. Increases in total dry
weights of starved oysters and oysters fed a 0.2% effective
ration resulted, therefore, from increases in ash content,
probably as a result of shell growth.

DISCUSSION

In bivalve growth experiments carried out by Langton
and McKay (1975) and Gallager and Mann (1981), ration
was not adjusted according to growth over the entire
experimental period and the animals were fed a constant
amount of food per individual. An important consequence
of maintaining a constant ration with rapidly growing
animals is that the weight-specific ration (expressed as a
percentage of oyster live weight in this study) decreases as
the animal grows (Figure 1 ). An example of large decreases
in weight-specific ration is evident in Experiment 6 of
Walne and Spencer (1974) in which a ration of Tetraselmis
suecica (Kylin) Butch, fed to Ostrea edulis Linne decreased
from 35 to 2% of oyster live weight over a 3-week period.
This occurred even though the authors attempted to com-
pensate for oyster growth by limited, but insufficient.

URBAN ET AL.

TABLE 2.

The relationship between the effective algal ration and the resulting growth
and gross growth efficiency of juveniles of Crassostrea virginica.

Increase in

Dry Wt of Algae

Percent Increase in

Initial Oyster

Oyster

Oyster Dry

Final Oyster

fed per

Oyster Live Wt/

Gross Growth

Effective Ration*

Live Wt

Organic Wt

Organic Dry

Experiment

Dry Wt of Algae

Efficiencyf

(X 100)

(mg)

Wt/Live Wt

(mg)

fed per Experiment

(GGE)

2.8

224.76

3,014.3

151.12

0.051

715.5

420

21.1

2.8

228.17

2,774.2

114.48

0.043

615.2

450

18.6

2.2

315.78

2,135.6

85.30

0.040

475.7

450

17.9

2.3

221.60

1,853.7

75.25

0.043

383.6

480

19.4

1.6

224.64

1,265.0

46.88

0.041

231.1

550

20.3

1.6

225.43

1,337.8

48.08

0.040

271.5

610

22.1

1.2

227.85

723.0

28.42

0.044

143.4

500

19.8

1.4

557.65

1,671.7

78.77

0.046

347.4

480

22.6

1.0

224.96

362.4

38.34

0.085$

82.2

440

46.6 %

1.1

556.17

1,098.8

46.07

0.042

224.3

490

20.5

0.9

222.00

490.4

15.50

0.043

77.7

630

19.9

0.7

559.33

628.5

26.30

0.042

133.4

470

19.7

0.6

316.69

344.3

12.34

0.039

65.0

530

19.0

0.5

224.91

126.8

0.13

0.040

24.0

370

0.5

0.2

228.22

47.0

- 4.22

0.039

11.2

420

-37.7

0.4

556.11

219.7

6.35

0.038

57.2

380

11.1

unfed

219.17

48.2*

5.86

0.037

—

unfed

220.73

22.7*

- 6.43

0.039

—

unfed

551.66

31.9*

- 2.58

0.038

—

*Effective percent ration = average weekly effective percent ration for a 3-week experiment (Figure 1), expressed as (mg dry wt algae per

mg live wt oyster) X 100.
fGross growth efficiency (GGE) = (increase in oyster dry organic weight/total dry weight of algae fed) X 100 for an experimental period of

3 weeks.
tThese values are anomalous and may have resulted from analytical error.
♦Increases in the live weight of fed animals were adjusted by subtracting the mean increase in the live weight of starved animals. This was

necessary to accurately determine gross growth efficiency (Winberg 1958).

weekly increases in ration. Clearly, if a constant weight-
specific ration is desired throughout a growth experiment,
frequent adjustments of ration in proportion to bivalve
growth are necessary. Such adjustments are especially
important in growth experiments with juvenile animals in
which weight-specific growth rates are high, and which
result in significant changes in weight-specific rations over
short periods of time, unless frequent ration adjustments
are made. Changes in weight-specific rations will be less
dramatic with large animals that have lower weight-specific
growth rates. Under certain conditions the use of photo-
electric devices to maintain constant algal concentrations
may be useful (Winter 1973).

Pruder et al. (1976, 1977), Epifanio and Ewart (1977),
and Epifanio (1979) attempted to determine the maximum
ration that could be ingested by bivalves under optimal
growth conditions where excess food was available. Under
those conditions, they assumed that the growth rate would
be greatest when the animal was fed as much food as it
could consume, i.e., a maximum ration (Epifanio and Ewart
1977). Because maximum ration is dependent on animal
weight (Navarro and Winter 1982), several ration formulae,
derived from measurements of the filtration rates of

(J
c
QJ

-t-

(/)

-20

-40

Effective
% Ration

Figure 3. Gross growth efficiencies of juveniles of C. virginica fed
different effective percent rations for a period of 3 weeks. GGE =
(increase in oyster organic dry wt/dry wt of algae fed) X 100.

Effect of Ration on Growth of Juvenile Oysters

Crassostrea virginica, have been described in an attempt to
predict the maximum ration on a weight-specific basis.
Pruder et al. ( 1 976) reported an empirically derived equation
relating oyster weight to a daily requirement of cells of a
mixture of Thalassiosira pseudonana and hochrysis galbana.
The equation Y = 5.3 W" 0,41 was derived on the basis of the
maximum filtration rates of both laboratory-reared juvenile
oysters and adult oysters from the field, where Y was the
daily ration of algal cells of a 50/50 mixture (by cell number)
of T. pseudonana and /. galbana X 10 8 per gram live weight
of oyster and W was the individual oyster live weight in
grams. Later, Pruder et al. (1977) repeated the work using
only laboratory-reared oysters and the equation was modi-
fied to Y = 8.2 W~°- 21 . The modification was required
because laboratory-reared oysters had a higher content of
organic material compared with wild oysters.

Epifanio and Ewart (1977) determined the maximum
dry weights of four algal species which could be filtered
from suspension by laboratory-reared oysters (C. virginica)
of 15 g live weight. They found that the maximum ration
cleared varied from 4 mg/g/day (0.4% ration) for T. pseu-
donana to 1 5 mg/g/day (1 .5% ration) for /. galbana. Using a
maximum ration of 4 mg/g/day and a value for the exponent
of -0.41 obtained from Pruder et al. (1976), Epifanio and
Ewart (1977) derived the equation R/W = 0.01 W 0A \
where R was the daily ration of algae in mg dry weight, and
W was the individual live weight of the animal in grams. In a
later paper, Epifanio (1979) adjusted the value of the
exponent to a theoretical value which was closer to the
empirical value of Pruder et al. (1977) and the formula
predicting ration size was given as R/W = 0.01 W -033 .

The growth of C. virginica fed on rations derived from
the formulae of Pruder, Epifanio, and co-workers has not
been studied experimentally. In Figure 4, the predicted
rations are compared with those of the present study. In
the first week, the 4.6% initial ration was lower than the
predicted ration of Epifanio and Ewart (1977), but higher
than the rations of Pruder et al. (1977) and Epifanio (1979).
As the animals grew, the predicted rations based on the
weight-specific equations decreased and in the second and
third week of the growth experiment, all were less than
the 4.6% initial ration used in the present study.

It was impossible to definitely determine which rations
given in Figure 4 would support the greatest oyster growth.
Juvenile oysters fed on the highest initial ration of 4.6%
in this study grew at a constant exponential growth rate
throughout the experimental period (Table 1, Figure 1),
and were not adversely affected by the high algal concen-
trations of the ration during the latter part of the experi-
mental period. The optimal ration for maximum growth of
juvenile oysters weighing 11 to 64 mg was, therefore,
probably greater than that predicted by the weight-specific
equations. Further study is necessary to test this hypothesis
with juvenile oysters weighing less than 1 g, because the
equations of Epifanio and Ewart (1977) and Pruder et al.

(1977) were derived from experiments using larger oysters
than those used in the present study.

LEGEND

A - Ep.(amo& Ewart (1977)

Mean oysier hve *l B " Epifanio (1979)

11 3 ma C ~ Thu paper

D - Pruder eial 11977)

Mean oyster hvewl Mean oysler live v

p _ i | 1 ?•<»! t , 627

—

0-1 1-2 2-3

Experimental period
(weeks)

Figure 4. A comparison of the initial percent rations used in the
present paper and initial percent rations derived from reported
equations for determining the maximum ration for Crassostrea
virginica. The initial weekly mean individual live weights of oyster
fed the 4.6% ration in the present study are indicated above each set
of bars. These weights were used to calculate initial percent rations.
The 8-part bar "C" indicates the eight rations used in the present
study (Table 1).

The relationship between ration size and gross growth
efficiency (Figure 3) is similar to that reported for Mytilus
edulis by Thompson and Bayne (1974) in that there was
initially a dramatic increase in gross growth efficiency to a
maximum value, followed by a slight decline with further
increases in ration. At still higher rations, gross growth
efficiency may decrease even more sharply, making it
important for commercial oyster culturists to balance the
cost savings of further improvements in growth rate with
the increased costs of decreased utilization of expensive
algal food. Comparisons between gross growth efficiencies
of M. edulis and those reported in this paper for C. virginica
are difficult because Thompson and Bayne (1974) used
larger animals and also expressed gross growth efficiency
in terms of tissue dry organic weight and not total organic
weight (i.e., they did not include the contribution of the
food to synthesis of the organic fraction of the shell).
Price et al. (1976) reported that 39% of the total organic
material of M. edulis (3.5 to 14.4 g live weight) was present
in the shell and that 72% was present in the shell of adults
of C. virginica (80.9 to 170 g live weight). For juveniles of
C. virginica (10 to 30 mg live weight), the proportion of
the total organic matter present in the shell is 33.8 ± 5.8%
(C. Langdon, University of Delaware, Lewes, DE, unpub-
lished data). Clearly, failure to take into account increases in

URBAN ET AL.

the organic content of the shell may result in considerable
underestimations of gross growth efficiencies (see Jorgensen
1976).

Based on measurements of the total increase in the
organic weight of juvenile oysters, Romberger and Epifanio
(1981) reported a maximum gross growth efficiency of
36% for C. virginica fed a 50/50 mixture (by cell volume)
of T. pseudonana and /. galbana at ration levels based on
the predicted rations of Epifanio and Ewart (1977). Their
maximum gross growth efficiency was, therefore, greater
than the highest efficiency found in this study of 22.6%
and may have resulted from differences in culture conditions.

In conclusion, the use of high-algal rations and high
concentrations of algae up to 500,000 cells mvT 1 need not
be detrimental to oyster growth or growth efficiency when

used in batch-feeding systems (Pruder and Greenhaugh
1978). The highest initial percentage ration tested in this
study of 4.6% was greater than those recommended for
oysters of the same size by the predictive equations discussed
above. Constant adjustments of ration are required to
compensate for increases in oyster weight during the course
of growth experiments. An initial daily ration of 4.6%,
which was equivalent to an effective daily ration of 2.8% per
week, supported good growth of juveniles of C. virginica
under the conditions of this study. Optimal rations for
maximum oyster growth will vary according to culture
conditions. Empirical growth studies, such as those described
here, are useful because they integrate culture conditions
with both the physiological and nutritional requirements of
oysters for maximum growth.

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bivalve larvae. Pruder, G. D., C. J. Langon & D. E. Conklin, eds.

Effect of Ration on Growth of Juvenile Oysters 57

Proceedings of the Second International Conference on Aqua- Winberg, G. G. 1958. Rate of Metabolism and Food Requirements

culture Nutrition: Biochemical and Physiological Approaches to of Fishes. Nanchor. Trudy belojussk. gos. Univ. V. I. Lenina.

Shellfish Nutrition. 1981 October 27-29. Rehoboth Beach, DE. (Translated from Russian by Fish. Res. Board Can. Trans!. Ser.

World Maricult. Soc. Spec. Publ. 2:272-291. No. 194, 1960.)

Widdows, J. 1978a. Physiological indices of stress in Mytilus edulis. Winter, J. E. 1973. The filtration rate of Mytilus edulis and its

/. Mar. Biol. Assoc. U.K. 58:125-142. dependence on algal concentration, measured by a continuous

. 1978b. Combined effects of body size, food concentration, automatic recording apparatus. Mar. Biol. (Berl.) 22:317-328.

and season on the physiology of Mytilus edulis. J. Mar. Biol. . 1974. Growth of Mytilus edulis using different types of

Assoc. U.K. 58:109-124. food. Ber. Dtsch. Wiss. Komm. Meeresforsch 23:360-375.

Journal of Shell fish Research, Vol. 3, No. 1, 59-64, 1983.

EFFECT OF DEPURATION SYSTEMS ON THE REDUCTION OF BACTERIOLOGICAL
INDICATORS IN CULTURED MUSSELS (MYTILUS EDULIS LINNAEUS)

AURORA LEDO, ENRIQUE GONZALEZ, JUAN L. BARJA
AND ALICIA E. TORANZO

Departamento de Microbiologia

Facultad de Biologia

Universidad de Santiago de Compostela

Spain

ABSTRACT Five bacteriological parameters (total coliforms, fecal coliforms, fecal streptococci, Escherichia coli, and
total viable count) were used to examine depuration of cultured mussels (Mytilus edulis Linnaeus) by two different systems,
one using chlorine as a disinfection agent for the water, and the other using untreated seawater. The most significant
difference in post-depuration levels between chlorinated and untreated seawater systems was obtained for fecal coliforms
(63.4 and 90.1% reduction, respectively), whereas reduction of the other bacteriological parameters were quite similar for
both depuration methods. Although there was a large decrease in the fecal streptococci (> 74%), high residual numbers
could be detected after depuration. From the identification of bacteria isolated from mussels, we found that the pathogens
Salmonella and Yersinia were not recovered in the depurated samples, even though the genera Citrobacter, Enterobacter,
and Escherichia coli were detected either before or after depuration. The drug-resistance patterns of the most representative
members of the enterobacteria isolated from mussels were also determined.

KEY WORDS: mussels, Mytilus edulis, shellfish depuration, pollution indicators, drug-resistance

INTRODUCTION

Since Dogson (1928) found that depuration was an
effective method for reducing the microbial flora of contam-
inated shellfish, this method has been adopted as the best
technique for reducing the potential risk of public health
hazards associated with the consumption of shellfish which
might have accumulated high levels of bacterial or viral
pathogens.

In Galician "rias" (Atlantic coast of northwestern Spain),
the production of cultured mussels (Mytilus edulis Linnaeus )
on rafts is a very important economic activity, reaching
200,000 metric tonnes in 1981. Approximately 50% of
this production is destined for daily consumption and
export, following depuration which is required by Spanish
regulations.

The depuration process is based on holding shellfish in
tanks containing seawater that has been sterilized by
physical or chemical means. The technology of depuration
has been well studied (Huntley and Hammerstrom 1971,
Neilson et al. 1978, Souness et al. 1979), and reviewed
(Furfari 1976, Fleet 1978). Most countries have chosen to
clean their shellfish in depuration plants rather than by
relaying in natural waterways. Ultraviolet irradiation,
ozonation, and chlorination are widely used to sterilize
seawater for depuration (Kelly 1961, Wood 1961, Anon.
1972); however, Reynolds (1956) showed that the process
could be simplified if depuration plants were located in
areas with light or no contamination. In the former cases,
the water sterilization step could be suppressed. Because of
the special geography of Galician n'as, it is possible to find
within 30 km (18 miles) depuration plants located in areas

without microbial contamination, as well as others, nearer
populated areas (on the middle upper part), that must use
disinfection agents for water treatment.

Our objective was to compare the reduction of bacterio-
logical indicators of pollution in cultured mussels which
were subjected to depuration systems that used either
chlorinated seawater or untreated seawater.

MATERIAL AND METHODS

The sampling area selected for this study is located in
northwestern Spain (Figure 1). Mussel samples were
collected from January to June 1982, from rafts located in
several shellfish-growing areas, and were treated in three
different depuration plants; two plants used chlorinated
seawater and the other used untreated seawater.

During the sampling period, the water salinity ranged
from 31.7 to 34.3 ppt and the temperature oscillated
between 13 and 19°C. Total coliform levels of the water
in the chlorine-treated systems ranged from 230 to 830 per
100 m2. The standard dose of chlorine for water treatment
was 3 ppm. Treated water was dechlorinated by an appro-
priate aeration period before the mussels were placed into
the shellfish tanks. In the untreated system, the detected
level of total coliforms was never higher than 9/100 mC. In
both the treated and untreated systems the depuration time
period was 48 hours.

Samples were taken twice a month before and after
depuration, transported to the laboratory in isotherm con-
tainers, and immediately processed. Each sample was
divided into two subsamples which were analyzed simul-
taneously. Mussels were shucked aseptically according to

Ledo et al.

Depuration of mussels by Two Different Systems

procedures recommended for shellfish by the American
Public Health Association (APHA 1970). One hundred grams
( 1 00 g) of shellfish meat without mantle fluid (corresponding
to six mussels) were weighed aseptically. After the addition
of 1% of peptone water, the mixture (1 :9 w/v) was homoge-
nized for 60 seconds in a sterile Waring blender. Each
homogenate was transferred into a sterile flask and used as
inoculum. Ten-fold serial dilutions of the homogenate were
inoculated in triplicate on plate-count agar (Difco) and
incubated at 37°C for 24 hours. After incubation, plates
were counted and the results were expressed as colony-
forming units (CFU) per gram.

Total coliforms were estimated by the standard most
probable number (MPN) method using three dilutions in
three tube replication of lactose broth (LB) (Difco). Tubes
were incubated at 35°C for 48 hours after which they were
examined for growth and gas production (APHA 1970).
Lactose broth tubes were reinoculated simultaneously into
brilliant-green lactose bile broth (BGLB) (Difco) and into
1% triptone water, then incubated in a water bath at 44.5 ±
0.2°C for the indol test.

Tubes showing growth and gas in BGLB were confirmed
as fecal coliforms (FC). The MPN of Escherichia coli was
determined from positive tubes for both tests, growth with
gas at 44.5 ± 0.2°C and indol production.

Fecal streptococci were determined by the MPN method
in azide dextrose broth (Difco) at 35°C. Positive tubes of
presumptive test were inoculated in ethylviolet-azide broth
(Difco) at 35 C. Tubes showing violet sediment were con-
sidered positives and the presence of fecal streptococci was
confirmed by streaking on KF-streptococcus agar (Difco).

Positive tubes from LB and BGLB of the MPN test were
streaked on Levine-eosin methylene blue agar (Difco) and
incubated at 37°C for 24 hours to isolate enterobacteria.
Colonies were picked randomly from the plates, subcultured
repeatedly to obtain pure cultures, and stored on agar slopes
under mineral oil at room temperature. The isolates were
subjected to taxonomic analysis using morphological,
physiological and biochemical tests according to the pro-
cedures of Edwards and Ewing (1972) and Bergey's Manual
(Buchanan and Gibbons 1974).

The drug-resistance patterns of the isolates were deter-
mined by the diffusion disk assay method of Bauer et al.
(1966) on Mueller-Hinton agar (Difco). The following anti-
biotics and concentrations were used: ampicillin (10 jug),
chloramphenicol (30 jug), erythromycin (15 /Jg), gentamicin
(10 jug), polymyxin B (300 units), nalidixic acid (30 jug),
kanamycin (30 jug), tetracycline (30 jug), and streptomycin
(10 jug).

RESULTS AND DISCUSSION

The results obtained in this study of depuration levels of
total coliforms (TC), fecal coliforms (FC), fecal strepto-
cocci (FS). Escherichia coli, and total viable count (TVC)
with the two systems used are shown in Table 1 and Figure 2.

Total viable counts*

61.5

Total coliformsf

30.2

Fecal coliformsf

63.4

Escherichia co//f

91.5

Fecal streptococcif

74.0

In general, only small differences were observed between
the two depuration systems. For the total viable count,
similar values were obtained. The TVC decreased by 10-fold
over the depuration time, but rarely went below values of
10 3 to 10 4 CFU/g of mussel. Similar results were found by
Lee and Pfeifer (1974) who worked with oysters depurated
by ultraviolet irradiated seawater and, as they indicated,
that reduction in bacterial count in shellfish could have been
due to the persistance of a stable population of micro-
organisms in the mussels. In addition, Thi Son and Fleet
(1980) obtained even lower reduction levels than ours in a
laboratory depuration system with artificially contaminated
oysters.

TABLE 1.

Comparison between the reduction levels of bacterial

pollution indicators in Mytilus edulis obtained

in two different depuration systems.

Percent Reduction in Systems Using

Bacterial Indicators Chlorinated Sea Water Untreated Sea Water

65.5
38.6
90.1
89.0
87.0

*Determined on plate-count agar medium at 37 C and expressed as

bacterial numbers per gram.
tDetermined by the most probable number (MPN) method and

expressed as MPN/ 100 g.

The most important different in the observed depuration
in chlorinated and untreated seawater systems was obtained
for FC, although in both methods most (about 90%) of
this bacterial flora was represented by E. coli. The high
depuration levels found for this organism agreed with the
the results obtained by Thi Son and Fleet (1980) who
attained depuration reductions greater than 97%.

Considering only the reduction rates for E. coli, we
found residual counts to be within the values allowed by
Spanish regulation (500 E. coli/9.) in both depuration
systems. If, however, we consider other regulations that
use the number of FC as the indicator for bacteriological
control, then the untreated seawater system appeared to
be the most efficient method (Table 1). The FC levels in
this system after depuration were below the recommended
wholesale level of < 230/100 g (Slalyj 1980) suggested by
the U.S. National Shellfish Sanitation Program for naturally
harvested shellfish.

Examination of bacteria isolated from mussels showed
that the genera Citwbacter, Enterobacter and Escherichia
coli were detected before and after depuration whereas
other pathogens or potential pathogens such as Salmonella
and Yersinia were not isolated from depurated samples of
mussels (Figure 3). The elimination of organisms such as

LEDO ET AL.

4_.

9
O

3-.

o 2_.

O
o

TVC

E.coli

*-/A

Chl Oc Chi Oc Chi Oc Chi Oc Oil Oc

Figure 2. Comparison between the reduction rates of bacteriological indicators obtained by the two different methods employed.

</)

_i
0.

s
<
en

<
t-
z

111

o
a.

□ BEFORE DEPURATION
■ AFTER DEPURATION

= O

c
o

D
CO

3
0}

o
o

Figure 3. Distribution of bacteria obtained from mussels before and after depuration.

Salmonella sp., Vibrio parahaemolvticus, and other patho- these bacteria were present in mussels before and after

gens during 48-hour depuration periods was also demon- depuration. This result supports the described higher survival

stratedby Metcalfet al. (1973) and Thi Son and Fleet(1980). of FS with respect to other bacteriological indicators in

Although the reduction levels obtained for FS were the marine environment (Cohen and Shuval 1972, Anson

similar in both systems (Table 1), very high numbers of and Ware 1974).

Depuration of Mussels by two Different systems

We determined the sensitivity of the enterobacteria
isolated from mussels to antibiotics and chemotherapeutic
agents; 77% of the strains displayed resistance to two or
more antibiotics. Table 2 shows the resistance patterns of
the most representative members of enterobacteria isolated:
E. coli, Citrobacter, and Enterobacter-Klebsiella group.
The percentage of E. coli strains resistant to tetracycline
was 44.5%, with the most frequent pattern being
erythromycin-tetracycline resistance. Most (90.8%) of the
Citrobacter strains were resistant to streptomycin, showing as
predominant resistance pattern erytromycin-streptomycin.
Of the isolates belonging to the Enterobacter-Klebsiella
group. 69.2% were resistant to ampicillin, with the
predominant pattern erythromycin-ampicillin.

Resistance to polymyxin and nalidixic acid was found
only in the genus Citrobacter, whereas resistance to chloram-
phenicol, gentamicin, and kanamycin was present only in
E. coli and Enterobacter-Klebsiella group strains, associated
with multi-resistant patterns.

It has been demonstrated that plasmids present in
enterobacteria codify drug resistance (Stewart and Kodit-
scheck 1980), as well as a variety of characteristics like
virulence (Elwell and Shipley 1980, Gemski et al. 1980,
Jones et al. 1982), enterotoxin production (Gyles et al.
1974, 1977; Mazaitis et al. 1981), and metabolic properties
such as urease production and citrate utilization (Gavini
et al. 1981), which could explain the relatively high number
of unidentified strains found in our study (Figure 3). Work
in progress indicates that these strains are multiplasmidic and
preliminary results have been presented (Barja et al. 1982).

TABLE 2.

Resistance patterns at two or more antibiotics in the most

representative members of enterobacteria

isolated from Mytilus edulis.

Bacterial Strains

Resistance Patterns*

Percentage

Escherichia coli

ETe

36.1

(36 strains)f

8.3

E Am

2.8

ESTe

2.8

ESC Am

2.8

E Te C Am

2.8

ESTeCKGm

2.8

Citrobacter

50.0

(22 strainslf

E Am

4.5

ESTe

18.2

E S Am

13.6

ESNa

4.5

ESPb

4.5

ETePb

4.5

En terobacter- Klebsiella

18.7

(16 strains)f

E Am

43.7

E Am Te

12.5

E S Te Am

6.5

E S Te C K Gm Am

6.5

*E, erythromycin; Te, tetracycline^, streptomycin; Am, ampicillin;

C, chloramphenicol; K, kanamycin; Gm, gentamicin; Na, nalidixic

acid;Pb, polymyxim.
fNumber of strains tested.

ACKNOWLEDGMENTS

The authors thank Dr. Francisco Lopez Capont (Dept.
Tecnologfa Pesquera, Facultad de Biologia, Universidad de
Santiago de Compostela. Spain) for sampling facilities.

REFERENCES CITED

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Anonymous. 1972. Use of ozone in seawater for cleansing shellfish.
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Anson, A. E. & G. C. Ware. 1974. Survey of distribution of bacterial
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Buchanan, R. E. & N. E. Gibbons. 1974. Bergey's Manual of Deter-
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Cohen, J. & I. J. Shuval. 1972. Conforms, fecal coliforms and fecal
streptococci as indicators of water pollution. Water Air Soil
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Dogson, R. W. 1928. Report on mussel purification. Fish. Invest.
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Edwards, P. R. & W. H. Ewing. 1972. Identification of Entero-

bacteriaceae. 3rd ed., Minneapolis, MN: Burgess Publishing Co.

362 pp.
Elwell, L. & P. L. Shipley. 1980. Plasmid irradiation factors associ-
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34:465-496.
Fleet, G. H. 1978. Oyster depuration: a review. Food Technol.

Aust. 30:444-454.
Furfari, S. A. 1976. Shellfish purification: a review of current

technology. FAO Tech. Conf. Aquaculture. FIR/AQ/Conf./

79/R.II. Kyoto, Japan.
Gavini, F., D. Izard, P. A. Tinel, B. Lefebvre & H. Leclerck. 1981.

Etude de taxonomique d'enterobacterie's appartenant ou

apparente'es a l'especie/T. coli. Can. J. Microbiol. 27:98-106.
Gemski, P., J. R. Lazere.T. Casey & J. A. Wohlhieter. 1980. Presence

of a virulence associated plasmid in Yersinia pseudotuberculosis.

Infect. Immun. 26:1044-1047.
Gyles, G. L., M. So & S. Falkow. 1974. The enterotoxin plasmids of

Escherichia coli. J. Infect. Dis. 130:40-49.
Gyles, G. L., S. Pachaudhuri & W. K. Mass. 1977. Naturally occurring

plasmid carrying genes for enterotoxin production and drug

resistance. Science 198:198-199.
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12:231-239.

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Jones, G. W., D. K. Kabert, D. M. Suinarich & H. J. Witfield. 1982.
Association of adhesive, invasive and virulent phenotypes of
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Kelly, C. B. 1961. Disinfection of seawater by UV radiation. Am. J.
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Lee, J. S. & D. Pfeifer. 1974. Influences of recovery media and incu-
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Maizaitis, J. A. & R. Mass. 1981. Structure of a naturally occurring
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Metcalf, T. G., B. Mullin. D. Eckerson, E. Moulton&E. Larkin. 1979.
Bioaccumulation and depuration of enteroviruses by the soft-
shelled clamAfva arenaria. Appl. Environ. Microbiol. 38:275-282.

Neilson, B. J., D. S. Haven, F. O. Perkins, R. Morales-Alamo & M. W.
Rhodes. 1978. Bacterial depuration by the American oyster
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Reynolds, N. 1956. A simplified system of mussel purification.

Fish. Invest. Sen II. Fish. G.B. Minist. Agric. Fish. Food 20(8):

18 pp.
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Amsterdam, The Netherlands: Elsevier Sci. Publ. Co.
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depuration of the Sydney rock oyster Crassostrea commercialis.

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Stewart, K. R. & L. Koditschek. 1980. Drug resistance transfer in

Escherichia coli in New York Bight sediment. Mar. Pollut. Bull.

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Thi Son, N. & G. H. Fleet, 1980. Behaviour of pathogenic bacteria

in the oyster, Crassostrea commercialis during depuration,

relaying and storage. Appl. Environ. Microbiol. 40:994-1002.
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Journal of Shellfish Research. Vol. 3. No. 1, 65-69. 1983.

DOCUMENTATION AND IMPLICATIONS OF RAPID SUCCESSIVE

GAMETOGENIC CYCLES AND BROODS IN THE SHIPWORM

L YRODUS FLORIDANUS ( BARTSCH )

(BIVALVIA, TEREDINIDAE)

C. B. CALLOWAY AND R. D. TURNER

Harvard University

Cambridge, Massachusetts 02138

ABSTRACT A pair (male and female) of the shipworm Lyrodus floridanus (Bartsch) was removed from the wood and
observed over a period of 39 days. The female of this short-term larviparious species broods its larvae in its gills to the
straight-hinge stage and then releases them en masse. Gametogenic cycles and brood periods were concurrent and regular,
averaging 6.12 (N = 4) and 5.02 (N = 5) days in length, respectively. Problems associated with observing gametogenic cycles
and brood periods in single animals, as well as the importance of such data in life-history studies, are discussed. Life history
data on L. floridanus support its removal from the synonymy of/,, pedicellatus and establish it as a distinct species.

KEY WORDS: Teredinidae, Lyrodus, brooding, gametogenic cycles, veliger larvae, spawning, reproductive cycles. Bivalvia

INTRODUCTION

Lyrodus floridanus (Bartsch), a species of wood-boring
bivalve, is found in Florida and probably throughout the
Caribbean. It is closely related to the common Californian,
but probably widely distributed, Lyrodus pedicellatus
(Quatrefages) and, generally, cannot be distinguished from
that species on the basis of shells and pallets (Turner 1966,
Turner and Johnson 1971). While studying the reproductive
biology of L. pedicellatus, a long-term brooder that releases
its larvae in the pediveliger stage, we found that specimens
from Florida differed by releasing their larvae in the straight-
hinge stage (i.e., they were short-term brooders). This was
first noted by Turner and Johnson (1971), but at that time
it was thought that under stressed conditions L. pedicellatus
might release straight-hinge larvae. We now realize that
L. floridanus is a distinct species with a reproductive pattern
like that of Teredo navalis Linnaeus. In both of these species,
eggs are spawned into the suprabranchial cavity and passed
into the water tubes of the gills where they develop to the
straight-hinge stage. They are then released en masse and
complete their development to the pediveliger stage in the
plankton.

To compare fecundities of different species, in this case,
L. pedicellatus and L. floridanus, it is necessary to know
the number and sizes of gametogenic cycles (oviparous and
brooding species) or broods (brooding species) for individual
specimens. Observations of this type were made using a
pair (male and female) of L. floridanus and form the basis
of this paper.

MATERIALS AND METHODS

Animals used in this study were obtained from collecting
panels exposed in the intracoastal waterway at Pompano
Beach, Florida, from 26 October 1978 to 26 February 1979.
Panels were hand-carried to Harvard University, Cambridge,

Massachusetts, on 27 February, and placed in an Instant
Ocean aquarium with natural sea water at 19 to 20 C and
32 ppt. They were dissected on the evening of 27 February
(day 1 ) and two uninjured specimens, one male and one
female, of Lyrodus floridanus (Bartsch), the predominant
species found in the panels, were placed in a finger bowl
with 200 m2 of 0.22-/im filtered sea water and maintained
in an illuminated incubator at 19 to 20°C. The water and
the bowl were changed daily to prevent the build up of
bacteria. Because some shipworms are capable of supple-
menting their diet of wood with phytoplankton (Dean and
Back 1979, Pechenik et al. 1979), the animals were fed
Isochrysis galbana, a naked flagellate, after each water
change at a final concentration of 4 X 10 4 cells/m£. Obser-
vations on the condition of the gonads and gills of the
female were made at each water change and often at shorter
intervals to determine the time of spawning and larval
release. Although spawning of the male was not observed
nor was any obvious change in size of the gonads evident,
sperm were seen attached to eggs aborted by the female.
When the experiment was terminated upon the death of the
female on day 39, gonadal smears of both animals were
examined and their sexes confirmed.

RESULTS

Shipworms are good animals for an observational study
of this type because the visceral mass, pericardium, gonads
and gills, which are located posteriorly to the shell, are
clearly visible through the translucent mantle (Figures 1-4).
Once the animal is removed from the wood, it is possible
to observe development of the gonads and growth of the
larvae without disturbing the animal. The gonads are
located between the pericardium and the wood-storing
caecum, and the genital ducts open into the suprabranchial
cavity posteriorly to the visceral ganglion (Figures 1-4).

Calloway and Turner

Figures 1 through 4. Lyrodus floridanus. Intact animal showing major anatomical features through the translucent mantle. (1) Left lateral
view of an adult female that is brooding straight-hinge larvae in the gill. The enlarged ovaries indicate that it is in the latter stages of a
gametogenic cycle (2.7X). (2) Enlargement of anterior end of animal in Figure 1. Note straight-hinge larvae in gills and the enlarged ovaries
(4.3X). (3) Left lateral view of an adult female that has recently released larvae (gills are empty). The greatly enlarged ovaries indicate that
spawning is imminent (2.7X). (4) Enlargement of anterior end of animal in Figure 3 (4X). Legend: A, auricle; F, foot;G, gill;GL, gill with
larvae; O, ovary; P, pallets; PC, pericardium; S, siphon ;SH, shell. Scale bar = 5 mm.

Immediately after spawning the lumina of the ovarian
follicles and tubes are empty and appear as clear mantle-
colored tissues arranged in a dendritic pattern on the surface
of the caecum. The first observable change in the ovaries as
gametogenesis proceeds is the appearance of oocytes in
the lumina of the follicles. As the number of oocytes
increases, the follicles enlarge, obscuring the dendritic
pattern, and the ovaries begin to turn white (Figures 1 and
2). Just before spawning, greatly enlarged white ovaries
completely cover the caecum laterally and dorsally and
extend posterodorsally to terminate at the opening of the
genital ducts (Figures 3 and 4).

Spawning is rapid, probably less than one hour in dura-
tion. At the conclusion of spawning the gonads are empty
and clear. The eggs pass from the suprabranchial chamber
into the water tubes of the gill, thereby turning the dorsal
portion of the gills white. As development progresses the
color of the gills change from white, when they contain
eggs, embryos, or trochophore larvae, to pale lilac as the
embryonic shell (prodissoconch I) forms, and then gradually
to a bright lilac as the prodissoconch II begins to develop

and the larvae reach the straight-hinge stage. [The terms
prodissoconch I and prodissoconch II are used in the sense
of Waller (1981).] As the prodissoconch II begins forming,
individual larval shells can be seen within the gill. Similar
to spawning, larval release is rapid, probably requiring less
than one hour. The larvae pass from the water tubes of the
gill to the suprabranchial cavity and are expelled from the
parent through the excurrent siphon. They develop to the
settlement stage, competent pediveligers, as planktotrophic
larvae.

One reproductive cycle, defined here as the time from
one spawning to the next, is divisible into two parts that are
readily observable by an examination of the gills. During
the brood period, the time from spawning until larval
release, the gills contain eggs, embroys, or larvae (Figures 1
and 2); during the empty period, the time from larval
release until spawning, the gills are empty (Figures 3 and 4).

Observation of the animals continued until the female
died on day 39. During this period, we observed four com-
plete and two incomplete gametogenic cycles as well as five
brood periods. The first gametogenic cycle was underway

Gametogenic cycles and Broods in the Shipworm

when the animal was removed from the wood and the last
cycle was in progress when the female died. Larvae from all
five broods appeared normal. Straight-hinge larvae from
brood 1 at the time of release measured 77.8 ± 1.4 jum long,
66.2 ± 1 .6 fini high, and had a hinge length of 43.7 ± 0.3 ^im
(N = 20). These measurements agree closely with the size
of larvae released from undisturbed animals living in wood
(79.4 ± 4.2 jum long, 70.0 ± 1.4 urn high, and a hinge line
of 47.4 ± 1.1 jum; N = 20). A small number of eggs was
expelled from the parent at each spawning. Eggs in the
germinal vesicle stage had a diameter of 52.0 ± 0.6 /im
(N = 20) and approximated the size of the eggs of Teredo
navalis (50 to 55 jum) reported by Culliney (1975).
Throughout the remainder of the brood period very few
larvae were released from the gills and these were usually
associated with mechanical disturbance caused by changing
the water and bowl.

Figure 5 is a diagrammatic representation of the gameto-
genic cycles and brood periods constructed from observa-
tions of the times of spawning and larval release. Times of
spawning and larval release are designated as the midpoints
between the times of successive observations (Figure 5). We

recognize that Figure 5 is a qualified representation of the
data. First, gametogenic cycles are considered to begin
directly after spawning. This is not necessarily so. Although
follicles appear empty at this time, gametogenesis could
have already begun. Conversely, a period may exist between
spawning and gametogenesis. Such a period would, however,
be short because oocytes are seen in the ovarian follicles
within one day after spawning. Second, the length of gameto-
genesis is unknown. Consequently, in Figure 5, gametogenic
cycles are drawn as straight lines. The ovary fills gradually
and empties rapidly. Third, the magnitudes of gametogenic
cycles and brood periods are not quantified. They are repre-
sented simply as the condition of the gonads and gills.
During our observations the size of the full gonads and gills
did not differ perceptibly from gametogenic cycle to
gametogenic cycle and from brood to brood. Therefore,
magnitudes of both the gametogenic cycles and brood
periods are diagrammed equally. It should be noted that the
gills were empty during gametogenic cycle 1. The probable
explanation for this is that, as so often happens when
animals are removed from the water for long periods of
time during transport to the laboratory, larvae are aborted

Full

O -r-

z
o
o

Emptv-

FulL

Z
O

□

z
o

Empty.

/ Gametogenic
Cycle 1

Gametogenic
Cycle 2

/ Gametogenic
/ Cycle 3

Gametogenic

. Cycle 4

Gametogenic

, Cycle 5

Brood 1

Brood 2

Brood 3

Brood 4

Brood 5

Spawn

Larval Release

2 *

5 10 15 20 25 30 35 40

TIME (IN DAYS)

Figure 5. Diagrammatic representation of gametogenic cycles and brood periods constructed from the times of spawning and larval release
observed in a single female Lyrodus floridanus. Spawning and larval release periods are figured as midpoints of successive observations.

Calloway and Turner

at the time the panel is put into the aquarium. There were
no larvae in the gills when the animal was dissected from
the wood but gametogenic cycle 1 was underway. The
greater length of this cycle possibly resulted from trauma
induced by the collecting and dissecting procedures.

It is apparent from Figure 5 that: (1 ) gametogenic cycles
are concurrent with brood periods so that the animals are
ripe at the time of larval release and spawning of the next
cohort occurs almost immediately, leaving only a short
period when the gills are empty; and (2) durations of the
gametogenic cycles and brood periods are regular, having
mean times of 6.12 ± 0.49 days (N = 4) and 5.02 ± 0.38
days (N = 5), respectively. Our observations of the brood
period of five days in Lyrodus floridamis maintained at 19
to 20°C are in close agreement with the report of a 5-day
brood period in Teredo navalis grown at 25 °C (Culliney
1975).

DISCUSSION

Breeding seasons of shipworms are largely based on
field collections or panel studies because breeding seasons
correspond roughly to dates of larval settlement (Schel tenia
and Truitt 1954, Nair and Saraswathy 1971). Characteris-
tically, larvae settle throughout the year in most tropical
marine areas and seasonally in high latitudes or areas of
varying salinity. Three major life-history patterns are known
for the Teredinidae: oviparous, short-term larviparous, and
long-term larviparous (Turner 1966, 1971; Turner and
Johnson 1971 ). We know the duration of the free-swimming
larval period and relative fecundities per brood for these
various life styles. Some estimates of numbers of eggs or
larvae released during a given reproductive cycle have been
published. For example, Sigerfoos (1908) estimated that a
large female of Teredo dilatata Stimpson (= Psiloteredo
megotara [Hanley] ), an oviparous species, releases 10 8 eggs
in a single spawning; Grave (1928) stated that a large speci-
men of Teredo navalis, a short-term brooder, produces 5 X
10 s to 10 6 eggs per spawning; and Karande et al. (1968)
reported that the brood of a 50-day old female of Teredo
furcifera von Martens, a long-term brooder, contained
7X 10 3 larvae.

Two vital life-history statistics are missing for all of these
species, i.e., the number and the size of broods and gameto-
genic cycles that occur during the life time of a given
individual. Without these data we cannot determine total
fecundity of an individual nor can we meaningfully com-
pare fecundities of species with different reproductive
patterns. The most direct way to obtain these data is to
observe single animals; however, in the Teredinidae this
type of study is not without problems. To observe indi-
vidual shipworms, we removed them from the wood and
could feed them only on phytoplankton. The animals were
undoubtedly stressed, but. nevertheless, the durations of
the gametogenic cycles and brood periods were typical of
those for Teredo navalis and probably for other short-term

larviparous species. If one could have only a single animal
per panel and could pair a male and a female in the same
aquarium the problem of stress would largely be eliminated.
It would then be possible to observe times of spawning in
oviparous species or larval release in larviparous species.
Unfortunately, in the case of larviparous species, only the
number of broods and the length of the reproductive cycle
could be determined because spawning could not be
observed. It is, of course, impossible to obtain data from
the same animal on both the total number of eggs or larvae
produced and the time course of gametogenesis, because
the latter would require histological examination. However,
the magnitude of each brood can be determined by
counting eggs spawned or larvae released. In larviparous
species, if it is assumed that no wholesale disintegration of
eggs or embryos occurs in the gills (we have seen no evidence
of this), then the number of eggs produced per gametogenic
cycle can be determined indirectly as the sum of aborted
embryos, aborted larvae, and released larvae.

Crisp and Davies (1955) have shown that if the values of
reproductive cycles and brood periods do not vary widely
about their means, then the fraction of the population
which is brooding is equal to the mean brood period divided
by the mean reproductive period. If the durations of the
brood and reproductive periods recorded for the single
Lyrodus floridamis which we observed are representative
of the population of L. floridamis in our test panels, then
87% of these animals would be brooding at a given time.
During the breeding season (which in Florida extends at
least from February through September and is probably
year around), we have often noted that the vast majority of
specimens dissected from the test panels were indeed
brooding.

This study, which began as a fortuitous observation,
dramatically illustrates another large gap in our knowledge
of the reproductive biology of the Teredinidae. A survey of
the marine invertebrate literature indicates that studies of
the reproduction of single animals with time are rare. The
paper on breeding of the barnacle Elminius modestus,
by Crisp and Davies (1955), is an excellent example of how
such investigations might be designed.

CONCLUSIONS

The documented rapid successive broods and gameto-
genic cycles in Lyrodus floridamis were unexpected and
explain why a large percentage of the animals in our
collecting panels contained eggs and larvae. These brood
periods and gametogenic cycles may also explain the
population explosions of short-term larviparous species that,
when introduced into a new area, may surpass native ovi-
parous species.

Turner (1966) considered L. floridanus a synonym of
L. pedicellatus mainly on the basis of shells and pallets of
preserved specimens. After observing living specimens in
Puerto Rico, Turner and Johnson (1971) suggested that

Gametogenic Cycles and broods in the shipworm

the pedicellatus-\ike Lyrodus, which released large numbers
of straight-hinge larvae, might be another species. Results
of the present research, combined with our unpublished
observations on morphological differences of the brood
pouches and of larvae, confirm the earlier suspicions of
Turner and Johnson (1971) that L. floridanus and/,, pedi-
cellatus are distinct species. The former broods its larvae
only to the straight-hinge stage and then releases them
en masse; the latter broods to the pediveliger stage, carries
several cohorts of larvae at different stages of development,
and releases only a few young at a time. Unfortunately,
young and nonbreeding specimens of these two species are
difficulty, if not impossible, to distinguish.

ACKNOWLEDGM ENTS

We are grateful to Ms. Paula Wagner for exposing and
retrieving the collecting panels, to Mr. Walter Baranowski
for drafting the figure, and to Drs. R. M. Woollacott and
J. A. Pechenik for reading the manuscript. The research was
supported by ONR Contract No. N00014-76-C-0281,
Nr 104-687 with Harvard University.

This paper was presented at the Nakhodka Symposium
on Physiology and Biochemistry of Adaptations in Marine
Animals in August 1979, as part of the 14th Pacific Science
Congress held at Khabarovsk, USSR.

REFERENCES CITED

Crisp, D. J. & P. A. Davies. 1955. Observations in vivo on the
breeding of Elminius modestus grown on glass slides. J. Mar.
Biol. Assoc. U.K. 34:357-380.

Culliney, J. L. 1975. Comparative larval development of the ship-
worms Bankia gouldi and Teredo navalis. Mar. Biol. (Berl.)
29:245-251.

Dean, R. C & G. G. Back. 1977. Suspension feeding on the ship-
worm Bankia gouldi (Mollusca; Bivalvia). Am. Zool. 17:948.

Grave, B. H. 1928. Natural history of shipworm, Teredo navalis, at
Woods Hole, Massachusetts. Biol. Bull. (Woods Hole) 55:260-282.

Karande, A. A., K. Balasubramanian & S. Prema. 1968. Development
of a laboratory method for bio-assay of candidate toxins against
teredid wood borers. Proc. Symp. Mollusca, Mar. Biol. Assoc.
India: p. 736-745.

Nair, N. B. & M. Saraswathy. 1971. The biology of wood-boring
teredinid molluscs. Adv. Mar. Biol. 9:335-509.

Pechenik, J. A., F. E. Perron & R. D. Turner. 1979. The role of
phytoplankton in the diets of adult and larval shipworms,

Lyrodus pedicellatus (Bivalvia: Teredinidae). Estuaries 2:58-60.
Scheltema, R. S. & R. V. Truitt. 1954. Ecological factors related

to the distribution of Bankia gouldi Bartsch in Chesapeake Bay.

Chesapeake Biol. Lab. Publ. 100:3-31.
Sigerfoos, C. P. 1908. Natural history, organization, and late

development of the Teredinidae, or ship-worms. Bull. U.S. Bur.

Fish. (1907)27:191-231.
Turner, R. D. 1966. A Survey and Illustrated Catalogue of the

Teredinidae (Mollusca: Bivalvia). Cambridge, MA: Harv. Univ.

Mus. Comp. Zool. 265 p.

. 1971. Australian shipworms. Aust. Nat. Hist. 17:139-145.

& A. C. Johnson. 1971. Biology of marine wood-boring

molluscs. Jones, E. B. G. & S. K. Eltringham, eds. Marine Borers,

Fungi, and Fouling Organisms of Wood. Paris: Organization of

Economic Cooperation and Development; p. 259-301.
Waller, T. R. 1981. Functional morphology and development of

veliger larvae of the European oyster, Ostrea edulis Linne.

Smithson. Contrib. Zool. 328:1-70.

Journal of Shellfish Research, Vol. 3, No. 1, 71-73, 1983.

RESEARCH NOTE

SETTLEMENT OF SPAT OF THE PURPLE-HINGE ROCK

SCALLOP HINNITES MULTIRUGOSUS (GALE)

ON ARTIFICIAL COLLECTORS

C. F. PHLEGER AND S. C. CARY

Department of Natural Science
San Diego State University
San Diego, California 92182

ABSTRACT Various artificial collectors were tested to obtain spat of the purple-hinge rock scallop Hinnites multirugosus
(Gale). These included plastic-mesh onion bags which were filled with nylon monofilament (gillnet), monofilament dipped
in cement, chaparral sticks, and a combination of sticks and empty scallop shells. The collectors were placed near a rock
scallop population in Mission Bay, San Diego, CA. The length of exposure and spatfall by season were also investigated.
Spat recruitment was greatest in gillnet collectors immersed for 3 to 4 months between late March and July. Up to 47 spat
of//, multirugosus (7 to 12 mm L) per gillnet bag were caught. Numerous spat of the blue musselMvf!7i« edulis Linne and
the wide-eared scallop Leptopecten latiauratus (Conrad) also settled in the gillnet collectors.

KEY WORDS: Rock scallop, Hinnites, spat collectors, spatfall, spat recruitment, aquaculture, mariculture.

INTRODUCTION

The purple-hinge rock scallop Hinnites multirugosus
(Gale) ranges from central Baja California to southern
Alaska and is common from the low-tide mark to 55 m
(Abbott 1974). Unlike the Atlantic bay scallop A rgopecten
irradians (Lamarck) and the Atlantic deep-sea scallop
Placopecten magellanicus (Gmelin), which are free-swimming
as adults, H. multirugosus cements itself to firm substrate
after a 6-month, free-swimming, juvenile (spat) stage. Like
the bay scallop it may temporarily attach by byssal threads.
The sessile nature of the adult has promoted considerable
aquaculture research with this species (Leighton and
Phleger, 1976, 1977, 1981; Cary et al. 1981). During this
study we addressed the problem of obtaining spat in
sufficient numbers for research or aquaculture development
and we employed experimental spat collectors to determine
the best settlement substrate, the appropriate immersion
time, and the period of greatest spatfall.

Spat of the Japanese scallop Patinopecten yessoensis( Jay )
can be collected with 1-mm mesh bags that contained mono-
filament gillnetting (Ito et al. 1975). Spat of .P. magellanicus
have been collected in 1.5-mm mesh onion bags which were
filled with monofilament gillnetting (Naidu et al. 1981).
Spat of the common European scallop Pecten maximus
(Linne) have been collected with Netlon®mesh envelopes
which contained nylon and plastic meshes and teased poly-
propylene rope (Brand et al. 1980). Thin monofilament
nylon has also been used as a substrate for settlement of
spat of the Iceland scallop Chlamys islandica (Miiller)
(Wallace 1981/82).

The molluscan taxonomy follows that of Abbott (1974)
for all but a few of the common bivalve names.

MATERIALS AND METHODS

Two principle types of spat collectors were used in
this study: (1) onion bags that contained 600 to 900 g of
loose, aquamarine monofilament (twine size #14, gill-
netting), and (2) plastic screen bags that were filled with
dry chaparral sticks. All of the bags were 42 X 75 cm and
1.0 to 1.5-mm mesh size. Spat bags were tied to concrete
pier pilings at a depth of 3 to 4 m and 3 m above the bottom
on the Ventura Bridge, Mission Bay, San Diego, CA, among
a large population of purple-hinge rock scallops. All deploy-
ment and retrieval of the spat bags were accomplished by
skin divers.

Scallops often attach to cement pilings. A series of spat
bags which contained gillnetting were partially coated with
Redi-Crete® cement to test its effectiveness as an attractant.
The cement dried and adhered readily to the monofilament
strands. Old rock scallop shells were included in a group of
screen bags (20 shells per bag) which also contained chaparral
sticks to act as an inducement for settling scallop spat.

Spat collectors were placed in the bay during the two
rock scallop spawning periods, late spring and late fall
(Jacobsen 1977). Fourteen gillnet bags (seven dipped in
cement) were placed in Mission Bay during December 1981,
and retrieved in March 1982. Twelve gillnet bags (without
cement) were placed in the same location and at the same
depth during March and June 1981. To determine the time
of spat settlement and seasonal growth rate, three bags
were retrieved at monthly intervals from June to September
1981. Screen- spat bags with chaparral sticks were placed in
the same Mission Bay location as the gillnet bags during
spring 1981. Eight stick-filled bags were placed in the bay
during April, May, and June 1981, and retrieved at 3-month

PHLEGER AND Cary

intervals. After retrieval, the spat bags were transferred to a
dock in Mission Bay and all newly settled scallops were
removed and counted. Because numerous invertebrates
attached to the gillnetting in addition to the rock scallops,
the gillnetting was repeatedly washed and shaken in sea
water in shallow plastic tubs to separate and recover the
spat and associated organisms.

RESULTS AND DISCUSSION

The spat of//, multirugosus were most abundant on the
gillnet collectors. Up to 47 spat occurred per bag and ranged
in length from 2 to 12 mm (mean lengths = 4 to 7 mm).
Plastic screen bags of sticks were much less effective in
attracting the spat. The total numbers of spat in the stick-
filled bags ranged from to 6, and spat lengths ranged from
3 to 9 mm (mean lengths = 5 to 7 mm). All 26 of the gillnet
spat collectors contained rock scallop spat, while only 6 of
the 24 stick-filled collectors from the same location con-
tained rock scallop spat. A Student's T-test showed no
significant difference at the p = 0.01 level between H.
multirugosus recruitment on cement-dipped gillnetting and
undipped gillnetting (Table 1). The addition of old scallop
shells to the stick-filled bags did not increase recruitment.
No rock scallop spat settled in two sets of four stick-filled
bags with and without old scallop shells which were set in
the bay at the same time and location. The success of
gillnetting versus other substrates may reflect its larger area
for attachment and subsequent growth of the scallop larvae
and spat.

Spat of the wide-eared (bay) scallop Leptopecten
latiauratus (Conrad) were invariably present in numbers of
up to 437 in gillnet collectors and up to 206 in stick-filled
collectors. Approximately 50% of the spat of/,, latiauratus
were dead (single shells or fragments), whereas all of the
spat of H. multirugosus were alive in the overwintered
gillnet collectors. Two bags with low numbers of spat (bags
2 and 3, without cement. Table 1) were torn open and

contained entangled fish hooks. Up to 100 living crabs
(Cancer spp.) were observed in the torn bags.

The time of spat settlement is important in the deploy-
ment of collectors for rock scallop spat. More spat attached
during the spring and early summer than during the pre-
ceding winter at the same location in Mission Bay. The
numbers of spat of//, multirugosus per bag ranged from 14
to 43 during three months in spring (24 March to 24 June
1982). The numbers of spat collected during the preceding
winter (Decmeber 1981 to March 1982) ranged from 2 to
24 (Table 1). In our previous study of recruitment of rock
scallops on the undersides of rock jetties in Mission Bay
during 1976 and 1977 (Leighton and Phleger 1981), we
also found small juveniles (3 to 10 mm, length) to be
abundant during late spring and early summer. Spatfall data
from the stick-filled bags showed that recruitment ceased
during May 1982. Eight stick-filled bags which were deployed
on 24 April 1982 and recovered on 24 June 1982 contained
16 spat (mean length = 6 mm). Spat length data suggest
that recruitment occurred only in March and April because
2-mm spat were about 2 months post-settlement. Spring
(March to April), therefore, appears to be the most appro-
priate time for deploying spat collectors for//, multirugosus
in southern California.

The fact that spat collectors, which were deployed during
spring and early summer, also contained large numbers of
spat of the blue mussel Mytilus edulis Linne (2,000 to
10,000 per bag) suggests that the rock scallop spatset may
have been much greater if there had not been such apparent
competition for setting space. Spat collectors that contained
gillnetting and that were over-wintered in the bay contained
only a few hundred blue mussel spat each. Other inverte-
brates whichwere recovered from the spat collectors included
free-living flatworms, juvenile gastropods, Hemphil's
swimming scallop Lima hemphilli Hertlein and Strong,
juveniles of Chione sp., pholad clams, polychaete scale and
serpulid worms, brachyuran crabs including Cancer sp.,

TABLE 1.

Results of trials with dipped and undipped spat collectors deployed in Mission Bay, San Diego, California

between December 1981 and March 1982.

Cement-Dipped M

onofilament Gillnetting

Monofilament Gillnetting Withoi

t Cement

Leptopecten

Hinnites

Percent of Total

Leptopecten

Hinnites

Percent of Total

Bag No.

latiauratus

multirugosus

(H. multirugosus)

Bag No.

latiauratus

multirugosus

/H. multirugosus)

134

7.5

115

4.3

151

10.6

12.5

172

14.0

44.4

113

3.5

164

3.7

114

1.8

175

5.7

3.3

6.5

206

2.9

16.3

Totals

980

6.6

Totals

673

7.3

Means

140

Means

RESEARCH NOTE

isopods. amphipods, arborescent bryozoans, juveniles of
the seastars Pisaster spp. and Asterina miniata (Brandt),
and the tunicate Ciona intestinalis (Linne). A few fish in
the genera Hyposoblennius and Girella were also recovered
from the spat collectors.

Spat collectors should not be deployed in the bay for
more than 4 months at a time. After 6 to 7 months of
immersion, numerous spat of H. multirugosus and almost
all spat of L. latiauratus were dead; we recovered mostly
single, empty, and many fragmented shells. The definitive
causes of spat mortality are unknown. Possible causes
include (1) anoxia detected in the spat collectors (H 2 S odor
and black sediment) which were held for 5 to 6 months,
and (2) crab (Cancer sp. and another unknown species) and
seastar (Pisaster spp.) predation. In some cases 25 to 100
crabs were recovered from infested spat collectors. We do
not know why anoxia and crab predation did not occur prior
to 5 or 6 months of exposure. The shells of/,, latiauratus
appear to be thinner than those of H. multirugosus and.
therefore, more susceptible to crab predation. Spat collectors

that were deployed for 3 to 4 months contained live spat
of//, multirugosus, but only empty or fragmented shells of
L. latiauratus.

This study indicated that spat collectors may represent a
practical method of obtaining large numbers of juveniles
(spat) of the purple-hinge rock scallop for an aquaculture
industry. Seasonability and total immersion time appear to
be the major factors that control the deployment and
effectiveness of spat collectors for//, multirugosus.

ACKNOWLEDGMENTS

We thank K. S. Naidu for providing 14 onion bag spat
collectors which contained gillnetting; D. L. Leighton pro-
vided advice and helped identify some of the invertebrates
in the collectors; and C. Wheatley, C. Papworth, and
N. Phleger provided field assistance. This research was
funded in part by NOAA, National Sea Grant College
Program, Department of Commerce, under Grant No.
NOAA-04-8-MOI-189, project R/A-44, and by the
California Resources Agency.

REFERENCES CITED

Abbott, R. T. 1974. American Seashells: Vie Marine Mollusca of

the Atlantic and Pacific Coasts of North America. (2nd ed.)

New York, NY: Van Nostrand ReinholdCo.
Brand, A. R., J. D. Paul & J. N. Hoogesteger. 1980. Spat settlement

of the scallop Chlamys opercularis (L.) and Pecten maximum

(L.) on artificial collectors. J. Mar. Biol. Assoc. U.K. 60:379-390.
Cary, S. C, D. L. Leighton & C. F. Phleger. 1981. Food and feeding

strategies in larval and early juvenile purple-hinge rock scallops

Hinnites multirugosus (Gale). J. World Maricul. Soc. 12(1):

156-169.
Ito, S., H. Kanno & K. Takahashi. 1975. Some problems on culture of

the scallop in Mutsu Bay.Bull.Mar. Biol. Stn.Asamushi 15:89-100.
Jacobsen, F. R. 1977. The reproductive cycle of the purple-hinge

rock scallop, Hinnites multirugosus (Gale) (Mollusca: Bivalvia).

San Diego, CA: San Diego State Univ. 75 p. Thesis.

Leighton, D. L. & C. F. Phleger. 1976. Preliminary studies on the
aquaculture potential of the Pacific Coast purple-hinge rock
scallop. Proc. World Maricul. Soc. 7:213 (abstract).

. 1977. The purple-hinge rock scallop: a new candidate

for marine aquaculture. Proc. World Maricul. Soc. 8:457-469.

. 1981. The suitability of the purple-hinge rock scallop

to marine aquaculture. San Diego State Univ., Center for Marine
Studies. Sea Grant Technical Rep. No. T-SCSGP001. 85 p.

Naidu, K. S., F. M. Cahill & D. B. Lewis. 1981. Relative efficacy of
two artificial substrates in the collection of sea scallops
{Placopecten magellanicus) spat. J. World Maricul. Soc. 12(2):
165-171.

Wallace, J. C. 1981/82. The culture of the Iceland scallop, Chlamys
islandica (O. F. Mu Her). I. Spat collection and growth during
the first year. Aquaculture 26:311-320.

Journal of Shellfish Research, Vol. 3, No. 1, 75-104, 1983.

ABSTRACTS OF TECHNICAL PAPERS

Presented at 1982 Annual Meeting

NATIONAL SHELLFISHERIES ASSOCIATION

Baltimore, Maryland

June 14-17, 1982

National Shellfisheries Association. Baltimore, Maryland Abstracts, 1982 Annual Meeting, June 14-17, 1982

CONTENTS

George R. Abbe

A Study of Blue Crab Populations in Chesapeake Bay in the Vicinity of the Calvert Cliffs

Nuclear Power Plant, 1968-1981 81

Philip Alatalo, Carl J. Berg, Jr. and Charles N. D Asaro

Reproduction and Development in the Lucinid Clam Codakia orbicularis Linne 81

Saved M. AH and G. D. Pruder

Effects of Inorganic Particles on the Growth of the Eastern Oyster Crassostrea

virginica (Gmelin) 81

Stand ish K. Allen

Applications of Flow Cytometry to Cytogenetic Studies in Bivalve Molluscs:

Measuring Changes in DNA Content 82

R. S. Appeldoorn, D. L. Ballantine and P. Chanley

Observations on the Growth and Survival of Laboratory-Reared Juvenile Conchs,

Strombus gigas and S. coastatus 82

Jenny A. Baglivo, George E. Lang and Diane J. Brousseau

A Simulation Study of a Stochastic Harvesting Model for Mya arenaria Linne 82

James M. Bishop and V. G. Burrell, Jr.

An Experimental Habitat Pot for Premolt Crab Capture 82

Jay A. Blundon and Victor S. Kennedy

Refuges from Blue Crab (Callinectes sapidus Rathbun) Predation for Infaunal

Bivalves in the Chesapeake Bay 83

Christopher F. Bonzek and Michael M. Burch

A Random Sample Survey to Estimate Blue Crab Catch in Maryland 83

Mark L. Bo t ton

What Determines the Vulnerability of Bivalve Prey to Horseshoe Crab Predation? 83

Neil Bourne

Clam Predation by Scoter Ducks in the Strait of Georgia, British Columbia 84

Diane J. Brousseau, Jenny A. Baglivo and George E. Lang

Determination of Settlement Rates in Shellfish Populations using Mya

arenaria Linne as a Model 84

M. Brouwer, D. Engel and J. Bonaventura

Heavy Metal Binding to Proteins of the Blue Crab Callinectes sapidus Rathbun 84

Carolyn Brown

The Role of Carbon Filtration in Culturing the American Oyster Crassostrea virginica (Gmelin) 85

John W. Brown, John J. Manzi, Harry Q. M. Clawson and Fred S. Stevens

Moving Out the Learning Curve: An Analysis of Nursery Operations for the Hard Clam

Mercenaria mercenaria (Linne) in South Carolina 85

Norman E. Buroker

A Survey of Allozyme Variation and Estimates of Genetic Similarity among Three Ostrea Species 85

Edwin W. Cake, Jr. and Vincent J. Smith

The Southern Oyster Drill: A Predator of Trapped Blue Crabs 85

Oral Capps, Jr.

Factors Affecting Dockside Prices for Hard Blue Crabs in Chesapeake Bay 86

Melbourne R. Carriker

Molluscan Shell Dissolution by Penetrating Eumetazoan Invertebrates: An Hypothesis

on the Chemical Mechanism based on Ultrastructure 86

Thomas P. Cathcart and Russell B. Brinsfield

Composting of Blue Crab Scrap: Problems and Solutions 86

Mark Chatry and R. J. Dugas

Optimum Salinity Regime for Oyster Production on Louisiana's State Seed Grounds 87

Timothy J. Cole

Gene Structures of Atlantic Coast Populations of the Blue Crab Callinectes sapidus Rathbun 87

78 Abstracts, 1982 Annual Meeting, June 14-17, 1982 National Shellfisheries Association, Baltimore, Maryland

CONTENTS (Continued)

John A. Commito

Naticid Snail Predation in New England: The Effects of Lunatia hews on the Population

Dynamics of Mya arenaria and Macoma balthica 87

/ D. Costlow and C. G. Bookhout

The Effects of Pollutants on Larval Development of the Blue Crab Callinectes sapidus Rathbun 87

L. Eugene Cronin

Analysis of Local Populations of the Blue Crab Callinectes sapidus Rathbun 88

Peter Daniel, Timothy Cole and Daniel Rittschof

Chemoreception and Life History of Stylochus ellipticus (Girard) 88

Ray C. Dintaman and J. F. Casey

Effect of Crab Pot Wire Treatment on Crab Pot Fouling in Chesapeake Bay, Maryland 88

Charles N. Dugas and M. Chatry

An Oyster Cultch Comparison: Clamshell versus Limestone 88

Elisa L. Elliot and Rita R. Colwell

Incidence of Pathogenic Bacteria in the Blue Crab Callinectes sapidus Rathbun and

the American Oyster Crassostrea virginica (Gmelin) 89

R. W. Elner and R. E. Lavoie

Predation on Spat of the American Oyster Crassostrea virginica (Gmelin) by the American

Lobster Homarus americanus Milne-Edwards, the Rock Crab Cancer irroratus (Say), and

the Mud Crab Neopanope sayi (Smith) 89

Charles E. Epifanio, C. C. Volenti and A. E. Pembroke

Seasonal Occurrence of the Larvae of Callinectes sapidus Rathbun in Delaware Bay 89

John W. Ewart and Melbourne R. Carriker

Characteristics of Fecal Ribbons from Juveniles of Crassostrea virginica (Gmelin) Fed

Phaeodactylum tricornuturn Bohlin With and Without the Addition of Silt: Preliminary Observations 90

Mary Jo Garreis and F. A. Pittman

Heavy Metal, Polychlorinated Biphenyl, and Pesticide Levels in Crassostrea virginica (Gmelin)

from Chesapeake Bay 90

Eugene L. Geiger, Russell B. Brinsfield and Fred W. Wheaton

Reduction of Dissolved Organics in Blue Crab Processing Plant Effluent 90

Reginald B. Gillmor and Herbert Hidu

Morphometric Patterns in Intertidal Bivalves 91

Joy G. Goodsell, R. A. Lutz, M. Castagna, and J. Kraeuter

Nonplanktotrophic Larval Development of Two Species of Continental Shelf Bivalves 91

Gregory L. Gruber

The Role of the Ventral Pedal Gland in Formation of an Egg Capsule by the Muricid

Gastropod Eupleura caudata etterae B. B. Baker 1951 : An Integrated Behavioral,

Morphological, and Histochemical Study 91

Nancy H. Hadley and John J. Manzi

Some Relationships Affecting Growth of Seed of the Hard Clam Mercenaria

mercenaria (Linne) in Raceways 92

Robert C.Hale

Mixed-Function-Oxygenase Enzyme Systems: Purpose and Possible Deleterious

Interactions with Organic Pollutants in the Blue Crab 92

Paul C. Hammerschmidt

Estimates of Juvenile Blue Crab Abundance in Texas Bays 92

Harold H. Haskin, Eric S. Wagner and Mitchell L. Tarnowski

The Surf Clam along the New Jersey Coast: Population Size, Recruitment, Growth Rates 93

Herbert Hidu, Standish Allen and Jon Stanley

Growth Performance of Cytochalazin-induced Triploids of American Oysters and

Soft-shell Clams 93

National Shellfisheries Association. Baltimore, Maryland Abstracts, 1982 Annual Meeting, June 14- 17, 1982 79

CONTENTS (Continued)

Anson H. Hines and Kathryn L. Comtois

Predation by Blue Crabs and Spot on Infaunal Communities in Central Chesapeake Bay 93

Lewis S. Incze

Oceanography of the Southeastern Bering Sea and Recruitment Processes in Two

Species of Tanner Crab 94

David F. Johnson

Species-Specific Differences in the Megalopal Distributions Related to Water Density Parameters 94

Todd C. Kamens

Mechanism of Shell Penetration by the Burrowing Barnacle Trypetesa lampas (Hancock),

(Cirripedia: Acrothoracia): An Ultrastructural Study 94

Jeffrey Kassner

Trace Metals in Shellfish and Growing Area Designation 94

VictorS. Kennedy, C. King and J. Blundon

Blue Crab Predation on Infaunal Bivalves: Relation to Optimal Foraging Hypotheses 95

George E. Krantz

Department of Natural Resources and University of Maryland Form New Cooperative

Shellfish Research Unit at Cnsfield 95

George E. Krantz, G. J. Baptist and D. W. Meritt

Three Innovative Techniques that Made Maryland Oyster Hatcheries Cost-Effective 95

Judith Krzynowek

Effect of Processing on Sterol and Fatty Acid Composition of Crabmeat 96

Andre C. Kvaternik and William D. DuPaul

Estimation of Standing Crop of Mercenaria mercenaria (Linne) in the James River,

Virginia, using Commercial Records 96

Mark D. Leslie and Robert S. Wilson

Effects of Light and Gravity upon the Motile Behavior of Trochophore Larvae of

Mercenaria mercenaria (Linne) 96

R. A. Lutz, J. G. Goodsell, M. Castagna and A. P. Stickney

Growth of Juveniles of Arctica islandica (Linne) in Experimental Containers 96

John J. Manzi, F. S. Stevens, Y. M. Bobo, V. G. Burrell, Jr. and Nancy H. Hadley

Size and Volume Relationships in Juveniles of Mercenaria mercenaria (Linne):

A Revision of Belding's Tables 97

/. R. McConaugha, D. R. Johnson and A. J. Provenzano

A Descriptive Model for the Conservation of Blue Crab Larvae in the Vicinity

of Chesapeake Bay 97

R. E. Miller

A Test of a Dart Tag for Juvenile Blue Crabs, Callinectes sapidus Rathbun 97

Robert J. Miller

Methods for Field Experiments with Baited Traps 97

K S. Naidu

A First Estimate of Indirect Fishing Mortality in the Iceland Scallop Chlamys islandica (Miiller) 98

Carter R. Newell

The Annual Glycogen Cycle in the Soft-Shell Clam Mya arenaria Linne from Maine 98

Carter R. Newell

The Effects of Sediment Type on Growth Rate and Shell Allometry in the Soft-
Shell Clam Mya arenaria Linne 98

Roger I. E. Newell and Stephen Jordan

Preferential Ingestion of Organic Material from the Consumed Ration by the

Oyster Crassostrea virginica (Gmelin) 98

Elliott A. Norse and Virginia Fox-Norse

Factors Limiting Abundance of Callinectes spp 98

80 Abstracts, 1982 Annual Meeting, June 14-17, 1982 National Shellfisheries Association, Baltimore, Maryland

CONTENTS (Continued)

Eugene J. Olmi, III and James M, Bishop

Total Width-Weight Relationships of the Blue Crab Callinectes sapidus Rathbun

from the Ashley River, South Carolina 99

A. J. Provenzano, J. M. McConaugha, and D. F. Johnson

Significance of the Neuston Layer in the Dispersal of Larvae of the Blue Crab

Callinectes sapidus Rathbun 99

Hauke K. Rask

Growth Enhancement of Mya arenaria Linne and Mercenaria mercenaria (Linne)

by Marine Macroalgae 99

Raymond J. Rhodes

Economic Considerations in Management of the Commercial Blue Crab Fishery 100

Daniel Rittscholf, R. Shepherd and M. Carriker

Chemical Ecology of Oyster Drills 100

/. W. Ropes, D. S. Jones, S. A. Murawski, F. M. Serchuk, and A. Jearld, Jr.

Documentation of Annual Growth Lines in the Ocean Quahog/1 rctica islandica Linne 100

Leonard A. Shabman and Tamara Vance

The Chesapeake Bay Blue Crab Fishery: Historical Trends and Emerging Issues 100

Terry M. Scholar

Management of the Blue Crab Fisheries in North Carolina: A Case History 101

Thomas M. Soniat and Sammy M. Ray

The Texas Oyster Study. I. Relationships between Available Food, Oyster

Composition, Condition, and Reproductive State 101

Thomas M. Soniat, Sammy M. Ray and Rezenat M. Darnell

The Texas Oyster Study. II. Models of Oyster Nutrition in the Natural Environment 101

S. Stiles, and ./. Choromanski

A Cytogenetic Method as a Tool for Assessing the Condition of Shellfish Larvae 102

Mark L. Swift and S. Lakshmanan

Isolation and Partial Characterization of a Malate Dehydrogenase from

Crassostrea virginica (Gmelin) 102

Edward R. Urban and G. D. Pruder

Comparison of the Growth of Crassostrea virginica (Gmelin) at Five Algal Ration Levels

with Specific Reference to Predictive Feeding Equations 102

WillardA. Van Engel

A Blue Crab Management Plan: Objectives and Responsibilities 102

W. F. Van Heukelem and S. D. Sulkin

The Behavioral Basis of Larval Dispersal and Recruitment in the

Blue Crab Callinectes Sapidus (Rathbun 103

Debra A . Weinheimer

Reproductive Periodicity of Busycon carica (Gmelin) in Waters off South Carolina 103

Elizabeth L. Wenner and Charles A. Wenner

Distribution, Size, and Sex Composition of Three Species of Callinectes in the

Coastal Habitat of the South Atlantic Bight 103

James C. Widman, Edwin W. Rhodes and P. A. Boyd

Nursery Culture of the Bay Scallop Argopecten irradians irradians (Lamarck)

in Suspended Mesh Enclosures 104

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

A STUDY OF BLUE CRAB POPULATIONS IN

CHESAPEAKE BAY IN THE VICINITY OF

THE CALVERT CLIFFS NUCLEAR

POWER PLANT, 1968-1982

GEORGE R. ABBE

Academy of Natural Sciences of

Philadelphia

Benedict Estuarine Research Laboratory

Benedict. Maryland 20612

Blue crab (Callinectes sapidus) population data were col-
lected from 1968 to 1981 to determine the effects of waste
heat from the Calvert Cliffs Nuclear Power Plant (CCNPP)
on abundance, size distribution, sex ratios, and seasonality.
Crabs were sampled using commercial crab pots of 2.5-cm
mesh set within (Plant Site) and outside (Kenwood Beach
and Rocky Point) the main thermal-effect area. Five pots
per station were fished 4 days/week during alternate weeks
from May through November. Crabs were sexed, measured,
and weighed by sex. In 14 years, a total of 10,552 pots
yielded 57,144 crabs (5.42/pot) of which 74.1% were legal
size ( > 127 mm carapace width) and 51.6% were male.
During 7 preoperational years (1968-74), crabs/pot aver-
aged 4.06 at Kenwood Beach (33.3%), 3.94 at Plant Site
(32.3%), and 4.18 at Rocky Point (34.3%). During 7 opera-
tional years (1975-81), crabs/pot averaged 6.24 at Kenwood
Beach (33.3%), 6.37 at Plant Site (34.0%), and 6.13 at
Rocky Point (32.7%). Increased catch during the operational
period was due to extreme abundance in 1981 when pots
averaged nearly 17 crabs. Data analyses revealed no signifi-
cant station differences other than a higher percentage of
males at Kenwood Beach than at Rocky Point (p=0.005).
There has also been a significant decrease in percent males
since 1968 (p < 0.001) which has occurred equally at all
stations. No effect of the CCNPP on crab populations was
evident from these studies.

REPRODUCTION AND DEVELOPMENT IN THE LUCINID
CLAM CODAKIA ORBICULARIS LINNE

PHILIP ALATALO 1 , CARL J. BERG 1
AND CHARLES N. D'ASARO 2

Marine Biological Laboratory
Woods Hole, Massachusetts 02543

Department of Biology
University of West Florida
Pensacola, Florida 32504

The tiger lucine Codakia orbicularis is a large edible clam
currently being investigated as a mariculture candidate in the
Bahamas Islands. Gonad development and spawning seasons
were assessed by monthly sampling of C. orbicularis from
Grand Bahama Island and Key Biscayne, Florida. Histological

examination of clams exceeding 20 mm in shell length
showed most of the populations sampled ripe between
April and November. Natural spawning probably occurs
from May to October.

Clams seldom respond to standard spawning techniques,
including physical and chemical stimuli. Artificial fertili-
zation by carefully stripping the gonads produced 15 to
20% viable embryos. Eggs are 108 to 112 nm in diameter
and are encased in a thick capsular membrane. Following
fertilization, the gastrula, trochophore, and early veliger
stages develop within the capsular membrane. Upon
hatching, the planktonic veliger ranges from 150 to 174 fxm
in shell length and develops to the pediveliger stage in
approximately 12 days. Metamorphosis occurs approxi-
mately 16 days after fertilization. Larval growth and
developmental features peculiar to C. orbicularis are
discussed.

EFFECTS OF INORGANIC PARTICLES ON THE
GROWTH OF THE EASTERN OYSTER
CRASSOSTREA VIRGINICA (GMELIN)

SAYED M. ALI AND G. D. PRUDER

College of Marine Studies
University of Delaware
Lewes, Delaware 19958

The effect of seven concentrations of inorganic particles
(oxidized silt from the Broadkill River) on the growth of
oysters (Crassostrea virginica) was studied at each of three
algal ration levels. In the absence of silt (zero concentration)
oyster growth was not significantly different between the
selected algal ration levels. At the lowest algal ration, the
addition of silt did not significantly affect oyster growth
rate; however, at the medium and high algal ration levels
oyster growth did increase with increasing silt concentra-
tion up to 25 mg/£. Above 25 mg/2, up to 150 mg/8. the
increased growth rate level was maintained showing neither
further enhancement nor any adverse effect on oyster
growth. The silt effect is discussed in terms of improved
delivery of food, growth factors, toxic metabolites, increased
digestability, resuspension of pseudofaeces, and increased
filtration and ingestion rates. Implications of the findings
for bivalve molluscan mariculture are suggested. The
increased growth rate could not be explained by any single
mechanism.

82 Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

APPLICATIONS OF FLOW CYTOMETRY TO CYTOGENETIC
STUDIES IN BIVALVE MOLLUSCS: MEASURING
CHANGES IN DNA CONTENT

STANDISH K. ALLEN, JR.

Marine Cooperative Fisheries
Research Unit
University of Maine
Orono. Maine 04469

Flow cytometry is a relatively new approach to cyto-
genetic studies in the biomedical field. This technique is of
considerable utility in other fields, especially in measuring
quantum shifts in DNA content. Diploid and triploid oysters
and clams were subjected to tissue disaggregation and nuclei
isolation techniques in an attempt to derive a suspended cell
population for analysis. Tissue disaggregation was shown to
be most effective and the principles of this method are des-
cribed. Nonlethal analysis of DNA content in individual
bivalves was also accomplished by sampling cells from hemo-
lymph sinuses. An apparent quantum duplication of DNA
between the sea scallop and bay scallop was demonstrated.
Recommendations for continued investigations using flow
cytometry are presented.

OBSERVATIONS ON THE GROWTH AND SURVIVAL OF

LABORATORY-REARED JUVENILE CONCHS,

STROMBUS GIGAS AND S. COSTATUS

R. S. APPELDOORN, D. L. BALLAN-
TINE AND P. CHANLEY

Department of Marine Sciences
University of Puerto Rico
Mayaguez, Puerto Rico 00708

A study of the culture and life history of the queen conch
Strombus gigas Linne in Puerto Rico has been underway
since 1981. Its objective is to develop suitable methods for
the large-scale culture of larvae of S. gigas and subsequent
release of juveniles to rebuild depleted natural stocks.
Although efforts have concentrated on S. gigas, larvae of the
closely related milk conch S. costalus Gmelin have also been
raised. Larvae were raised from eggs collected from the field.
The larval period was variable with settlement commencing
from 12 to 19 (x = 15.6) days after hatching. Length at
metamorphosis varied from 1.2 to 1.8 mm with a mode
between 1.4 and 1.5 mm. Sets of over 1,000 juveniles were
achieved with survival ranging from 4 to 7% from hatching to
a postmetamorphosis size of 3 to 5 mm. After metamor-
phosis growth increased noticeably. Initial postmetamorpho-
sis growth was 0.2 mm/day, but the rate of growth continued
to increase reaching a mean of 4 mm/day through the first
200 days. Feeding experiments of juveniles indicated that
the macroalga SpyriJia filamentosa (Wulfen) was preferred.

Pilot experiments involving the release of small (25 to
50 mm) tagged juveniles permitted the testing of suitable
mark and recapture methods and the collection of prelimin-
ary observations of juvenile behavior. These observations
indicated that mortality was initially high but dropped over

time. Dispersal has been slow and random. Observed growth
was slow, probably caused by the large amount of time
spent buried and hence inactive.

A SIMULATION STUDY OF A STOCHASTIC
MODEL FOR MY A ARENARIA

JENNY A. BAGLIVO 1 , GEORGE E.
LANG 2 AND DIANE J. BROUSSEAU 3

Department of Mathematics
Fairfield University
Fairfield, Connecticut 06430 and
Department of Biostatistics
Sloan-Kettering Institute
New York, New York 10021
2 Department of Mathematics
Fairfield University
Fairfield, Connecticut 06430

Department of Biology
Fairfield University
Fairfield, Connecticut 06430

Field data presented by Brousseau (1978, 1979) provided
estimates of age-specific fecundity and survival for the soft-
shell clam Mya arenaria. We have used these values in a
Leslie population model (1945, 1948) to estimate an equili-
brium settlement rate for clams in the first age class (Brous-
seau et al., in press). Settlement rates are highly variable in
nature, however, and the modelling efforts incorporate this
phenomenon!. An optimal harvesting strategy based upon
the Leslie model was published by Rorres and Fair (1975).
We have designed simulation studies which adapt their pro-
cedure as well as other similar procedures to a stochastic
environment and applied these strategies using the Mya
model. Preliminary results show that these methods do not
over exploit the population; however, they may be too
conservative.

AN EXPERIMENTAL HABITAT POT FOR
PREMOLT CRAB CAPTURE

JAMES M. BISHOP AND
V.G. BURRELL,JR.

Marine Resources Research Institute

South Carolina Wildlife and Marine

Resources Department

P.O. Box 12559

Charleston, South Carolina 29412

Three years of testing premolt (peeler) crab capturing
devices showed unbaited habitat pots to be a potential har-
vest gear in South Carolina estuaries. Two and one-half-
centimeter mesh wire was used for pot construction, and
pot design was similar to that for baited hard crab pots.
Tests were conducted 4 consecutive days/week in the Ashley

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

River from mid-April through mid-November, 1979, and
daily in the Wando River from April through June, 1980
and 1981. Primary objectives were to increase pot efficacy
and reduce pot construction cost and labor.

Results showed that plastic flagging tape interwoven
among the wire mesh did not increase catch rates: pots with
and without tape averaged 0.7 peeler/gear-day (one pot
with a soak time of 24 h). Two large entrance pots (61 X
61 X 45 cm) outfished 4 small entrance pots (61 X 61 X
30 cm) by 1.6 vs. 1.3 peelers/gear-day, respectively. Pots
fished in shallow subtidal mudflats captured a mean of 1 .7
peelers/gear-day whereas those in deep water ( > 3 m) cap-
tured only 0.7 peeler/gear-day. Highest capture rates were
obtained in June during each year. A maximum of 3.5
peelers/gear-day was obtained when large habitat pots were
fished on shallow water mudflats in June. Male peelers
accounted for 63% of 1,832 peelers caught in habitat pots
during 1981. Habitat pots require no bait and offer crabbers
a method of harvesting peelers in relatively consistent num-
bers throughout the shedding season.

REFUGES FROM BLUE CRAB (CALLINECTES SAPIDUS

RATHBUN) PREDATION FOR INFAUNAL

BIVALVES IN THE CHESAPEAKE BAY

JAY A. BLUNDON 1 AND VICTOR S.
KENNEDY 2

Department of Zoology
University of Maryland
College Park, Maryland 20742

Horn Point Environmental Laboratories
University of Maryland
Cambridge, Maryland 21613

Direct measurements of valve strength of various sizes of
Mya arenaria Linne, Macoma balthica (LinneV, Macoma
mitchelli Dall, and Mulinia lateralis (Say) compared to
measurements of blue crab chelae grip strength suggest that
the shells of these infaunal bivalves confer no resistance to
crushing by blue crabs. Also, blue crabs readily crushed
these species in the laboratory.

Possible refuges from predation afforded to theseinfaunal
bivalves were investigated. Bivalve size, depth of burrowing,
and density were measured in the field throughout spring
and summer 1981 . This survey, in conjunction with labora-
tory feeding experiments that offered M. arenaria burrowed
at various sediment depths to blue crabs, suggested that M.
arenaria and M. balthica obtain refuge from blue crab preda-
tion at deeper sediment depths. Bivalves burrowed beneath
an artificial submerged aquatic vegetation structure also
gained additional protection. These refuges, however, were
not absolute, but only relative to infauna burrowed less
deeply or in bare sand (mud) environments. Yearly sampling

of bivalve infauna in the Choptank River, Chesapeake Bay,
suggested thatM mitchelli and M. lateralis are able to persist
despite predation due to their high reproductive output.

A RANDOM SAMPLE SURVEY TO ESTIMATE
BLUE CRAB CATCH IN MARYLAND

CHRISTOPHER F. BONZEK AND
MICHAEL M. BURCH

Maryland Department of Natural
Resources, Tidewater Administration
C-2 Tawes State Office Building
Annapolis, Maryland 21401

In June 1981 the Maryland Department of Natural
Resources (MDNR) began operating a new system to esti-
mate the catch of blue crabs (Callinectes sapidus Rathbun) in
Maryland waters. The basis of the system is a stratified, ran-
dom sampling design developed by the Martin Marietta
Corporation, which allows MDNR to reliably estimate total
crab catch in Maryland by asking only a small fraction of all
crabbers to report their catch each month. This method
produced a total annual harvest estimate in 1981 of 29.5 X
10 6 kg (65 X 10 6 lb) live weight, nearly twice the highest
estimate produced under past systems. The estimate is based
on standard statistical techniques, and takes into account
the previously ignored factors of non-reporting by some
crabbers and the non-commercial catch. Estimates of fisher-
man effort are produced concurrently.

WHAT DETERMINES THE VULNERABILITY OF BIVALVE
PREY TO HORSESHOE CRAB PREDATION?

MARK L. BOTTON

Department of Zoology

Rutgers University

P.O. Box 1059

Piscataway, New Jersey 08854

Adult horseshoe crabs, Limulus polyphemus (L.), were
offered combinations of different size and species of bivalve
prey in a large aquarium. Gemma gemma (Totten)
(Veneridae), a small, thick shelled species, was avoided
when larger, thinner shelled clams such as Mulinia lateralis
(Say) (Mactridae) or Mya arenaria Linne (Myidae) were
available. Crabs did not differentiate between M. lateralis
and M. arenaria of comparable size; however, there was a
preference for M. lateralis over hard-shell clams, Mercenaria
mercenaria (Linne) (Veneridae), of equal size. Large individ-
uals of M. lateralis, > 10-mm shell length, were preferred
over smaller individuals of M. lateralis. Thus, both shell
length and shell thickness appear to influence the preference
of horseshoe crabs for bivalve prey. The largest available prey
species offered to L. polyphemus was Spisula solidissima

84 Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

(Dillwyn) (Mactridae); clams up to 45-mm shell length were
successfully opened. The method of consuming these
bivalves differed from the manner in which smaller prey
were handled, and is illustrated.

CLAM PREDATION BY SCOTER DUCKS IN THE
STRAIT OF GEORGIA, BRITISH COLUMBIA

NEIL BOURNE

Fisheries and Oceans,
Pacific Biological Station,
Nanaimo, B.C.. Canada V9R 5K6

Collections of three species of wintering scoter ducks,
the white-winged scoter, Melanitta deglandi (Bonaparte),
the surf scoter, M. perspicillata (Linnaeus), and the black
scoter, Oidemia nigra (Linnaeus), were made at two clam
beaches in southern British Columbia. Analyses of the crop
and gizzard contents showed that these ducks were feeding
primarily in the intertidal beach area. Molluscs, particularly
bivalves, were the most important food items in the diet.
The commercially important littleneck and Manila clams,
Protothaca staminea (Conrad) and Tapes philippinarum
(Adams and Reeve), respectively.comprised about two thirds
of the gut contents of the scoters. Scoters are important
clam predators in southern British Columbia; it was esti-
mated that a wintering flock of 200 scoters could remove 5
to 14.5 metric tons of littleneck and/or Manila clams from
these two beaches in a 6-mo period.

DETERMINATION OF SETTLEMENT RATES IN SHELLFISH
POPULATIONS USING MY A ARENARIA LINNE' AS A MODEL

DIANE J. BROUSSEAU 1 , JENNY A.
BAGLIVO 2 AND GEORGE E. LANG 3

Department of Biology
Fairfield University
Fairfield, Connecticut 06430

Department of Mathematics
Fairfield University
Fairfield, Connecticut 06430 and
Sloan-Kettering Institute
New York, New York 10021

Department of Mathematics
Fairfield University
Fairfield, Connecticut 06430

Egg loss, larval recruitment, and early post-larval mortal-
ity are often limiting factors in the establishment and main-
tenance of shellfish stocks; therefore, it is of interest to
ecologists to be able to make estimates of settlement rates
in such populations. This paper describes an indirect method
for estimating mortality rates during settlement in shellfish
populations for which demographic parameters (age-specific
fecundity and survivorship) are available. The equilibrium

settlement rate for a population olMya arenaria from Glou-
cester, MA, was calculated using the Leslie matrix. Empiri-
cally derived demographic parameters indicate that the
theroretical settlement rate required to maintain a steady
state population is 0.001462% or one egg out of approxi-
mately 68,400 surviving to a size of 2 mm.

HEAVY METAL BINDING TO PROTEINS

OF THE BLUE CRAB CALLINECTES

SAPIDUS RATHBUN

M. BROUWER, D. ENGEL AND
J. BONAVENTURA

Marine Biomedical Center

Duke Univeristy Marine Laboratory

and NMFS Southeast Fisheries

Center Laboratory

Beaufort, North Carolina 28516

Hemocyanin is the large, extracellular oxygen transport-
ing protein found in the hemolymph of the blue crab. The
oxygen-binding site consists of a binuclear copper center. In
addition to copper, blue crab hemocyanin invariably con-
tains a small amount of tightly bound zinc (approximately
0.2 atom of zinc per oxygen-binding site). This observation,
together with the fact that hemocyanins act at the interface
between the organism and its environment, prompted us to
investigate a possible role of these respiratory proteins in
trace metal transport or toxicity in the blue crab. In vitro
studies revealed that blue crab hemocyanin can indeed bind
a variety of heavy metal ions, all with very high affinities
(18 mercury, 14 cadmium, and 24 zinc ions per oxygen-
binding site). The interaction of cadmium and zinc ions with
blue crab hemocyanin increases its oxygen affinity ; mercuric
ions have an opposite effect. All three heavy metal ions
reduce the degree of cooperativity in oxygen binding. Cad-
mium and zinc ions were found to substitute for calcium,
which is a natural modulator of blue crab hemocyanin
function.

In vivo exposure of blue crabs to cadmium dissolved in a
flowing seawater system at 0.1 ppm or to cadmium-ladened
oysters did not result in measurable elevated levels of
cadmium in the hemolymph. The sites of cadmium accumula-
tion varied depending on the method of exposure. Seawater-
exposed crabs accumulated most of the cadmium in the
gills; the ions were bound to a low molecular-weight protein
(MW~ 10,000). This protein was purified by gel-permeation
chromatography and ion-exchange chromatography. Cad-
mium was the only metal associated with the purified
protein. Crabs exposed to cadmium-ladened oysters accumu-
lated most of the cadmium in the hepatopancreas, where it
was associated with a low molecular-weight cadmium/zinc-
binding protein. Ion-exchange chromatography showed the
gill and hepatopancreas proteins to be different, suggesting
that these proteins, which are presumably involved in trace
metal detoxification, are tissue specific.

National Shellfisheries Association, Baltimore, Maryland

Abstracts. 1982 Annual Meeting, June 14-17, 1982

THE ROLE OF CARBON FILTRATION IN

CULTURING THE AMERICAN OYSTER

CRASSOSTREA VIRGINICA

CAROLYN BROWN

National Marine Fisheries Senice,
Northeast Fisheries Center,
Milford Laboratory,
Milford, Connecticut 06460

Embryos and larvae of the American oyster Crassostrea
virginica (Gmelin) were reared in two types of "disinfected"
seawater. One type was filtered through two 10-/im orlon
filters and UV-irradiated; the second type was subjected to
the same treatments, except that an additional filtration
process through a carbon cartridge was inserted prior to the
UV irradiation step. The study compared embryonic devel-
opment of the 2-day-old larval stage, as well as survival and
growth of larvae to metamorphosis in the two types of
treated seawater. Data indicated that the percentage of live-
normal development was significantly greater in seawater
subjected to carbon filtration than in seawater without this
added treatment. Other data suggested success in rearing
oyster larvae to metamorphosis using carbon filtration only
when the larval cultures were changed daily. Seawater treat-
ment is but one aspect of the prevention regimen to be fol-
lowed. Sound sanitary practices also are described to reduce
the frequency of disease outbreaks in hatcheries.

MOVING OUT THE LEARNING CURVE: AN ANALYSIS OF

NURSERY OPERATIONS FOR THE HARD CLAM

MERCENARIA MERCENARIA (LINNE')

IN SOUTH CAROLINA

JOHN W. BROWN 1 , JOHN J. MANZI 2 ,
HARRY Q. M. CLAWSON 3 AND
FRED S. STEVENS 4

1 South Carolina Sea Grant Consortium,
Charleston, South Carolina 29412

Marine Resources Research Institute
Charleston, South Carolina 29412
3 'Trident Sea farms Co., 18 Broad St.
Charleston, South Carolina 29401

Marine Resources Research Institute
Charleston, South Carolina 29412

Trident Seafarms (a private corporation) and the State of
South Carolina (SC Wildlife and Marine Resources Depart-
ment) entered into a cooperative research agreement for the
commercial production of hard clams in 1980. The SC Sea
Grant Consortium provided partial funding for the scientific
research and some staff time for the economic analysis of
the first 15 months of nursery operation. Detailed cost and
production analysis are provided, along with a description
of the evolution of the nursery production protocols and of
the nursery design. During the period from September 1980

to December 1981, 19,733,000 seed clams were imported
into the nursery; of these 13,008,000 remained in the nursery
at the end of the year, 3,402,000 were planted in the field
with 14,700 returned to the nursery. The apparent mortality
was 3,337,700 clams during the 15 months. This 16.9%
mortality is misleading because of the rapidly increasing
number of clams in the nursery over the period of the
analysis. Beginning with the correction for mortality, a
detailed budget analysis is given and linear programming is
employed to determine optimal importation strategies.

A SURVEY OF ALLOZYME VARIATION AND

ESTIMATES OF GENETIC SIMILARITY

AMONG THREE OSTREA SPECIES

NORMAN E. BUROKER

Bureau of Biological Research, Rutgers,
The State University of New Jersey
Piscataway, New Jersey 08854

Three nonsibling Ostrea species (i.e., O. edulis Linne,
O. lurida Carpenter. andO. pennollis Sowerby) were studied
by horizontal protein electrophoresis with relation to levels
of genetic variation and similarity. The percentages of poly-
morphic loci per species were estimated as 27.6, 37.0, and
52.0 for O. edulis, O. lurida, and O. permollis, respectively,
based on an examination of 25 to 29 structural loci. The
mean observed heterozygosities per individual were esti-
mated as 9, 16, and 1 5% for O. edulis, O. lurida, and O. per-
mollis, respectively. A pairwise comparison of loci was made
between species which indicated that approximately 17% of
the loci studies were genetically identical while 55% had no
genetic similarity. The mean genetic identity across all loci
among the three species was estimated as 24.5%. Finally,
there seemed to be a correlation between the dispersal time
of the planktonic larvae and the levels of genetic variation
found within these nonsibling Ostrea species.

THE SOUTHERN OYSTER DRILL: A PREDATOR
OF TRAPPED BLUE CRABS

EDWIN W. CAKE, JR. 1 AND
VINCENT J. SMITH 2

Oyster Biology Section,
Gulf Coast Research Laboratory
Ocean Springs, Mississippi 39564
2 Route 3, Box F-52
Ocean Springs, Mississippi 39564

Southern oyster drills (Thais haemastoma floridana
[Conrad] ) are reported for the first time to attack and kill
mature blue crabs (Callinectes sapidus Rathbun) in commer-
cial crab pots. Trapped blue crabs were attacked by as many
as 54 drills of up to 80 mm in shell height. All affected crabs
were either ovigerous or recently spent females, and all
were simultaneously infested with the symbiotic acorn

Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

barnacle Chelonibia panda (Ranzani). Entry portals for the
proboscis of feeding drills included: (1) open skeletal wounds
caused by other trapped crabs, (2) internal skeletal openings
between the branchial chamber and the infrabranchial sinuses
at the bases of the gills, (3) stumps of autotomized pereio-
pods, and (4) holes rasped in the exoskeleton by the snails'
radulae. The attacks were attributed to at least two factors:
the presence of large numbers of drills in the crab harvest
area in the vicinity of Mississippi's offshore barrier islands,
and the opportunistic feeding behavior of the drills,
especially when confined with trapped crabs. Moribund
and/or dead crabs also attracted another carnivorous snail,
the cancellate cantharus, Cantharus cancellarius (Conrad).

FACTORS AFFECTING DOCKSIDE PRICES FOR
HARD BLUE CRABS IN CHESAPEAKE BAY

ORAL CAPPS, JR.

Department of Agricultural Economics
Virginia Polytechnic Institute and State
University, Blacksburg, Virginia 24061

The nature and the magnitude of selected factors hypo-
thesized to influence the ex-vessel price of hard blue crabs
in Chesapeake Bay were investigated. The data base used
consisted of monthly observations for the period January
1973 to June 1980. Seasonality, landings of hard blue crabs
in Chesapeake Bay, and the wholesale price of hard blue
crabs had significant impacts on the ex-vessel price. Landings
of hard blue crabs in the south Atlantic and the Gulf were
not statistically significant in influencing the ex-vessel price
of hard blue crabs in Chesapeake Bay. On the basis of the
estimated flexibility coefficients, total revenue to harvesters
could be incremented by increasing landings in Chesapeake
Bay throughout each season of the year.

MOLLUSCAN SHELL DISSOLUTION BY PENETRATING

EUMETAZOAN INVERTEBRATES: AN HYPOTHESIS

ON THE CHEMICAL MECHANISM BASED ON

ULTRASTRUCTURE

MELBOURNE R. CARRIKER

College of Marine Studies
University of Delaware
Lewes, Delaware 1 9958

Of the 27 eumetazoan invertebrate phyla generally recog-
nized, at least 8 widely separated ones are known to contain
shell penetrating species (burrowers or borers): Platyhel-
minthes, Bryozoa, Sipunculoidea, Phoronida, Annelida,
Arthropoda, Brachiopoda, and Mollusca. The pattern of
molluscan shell dissolution is similar at the ultrastructural
level in species of four phyla that have been studied:

polychaete Polydora websteri Hartman (Zottoli and Carriker
1974), barnacle Trypetesa lampas (Hancock) (Todd 1981).
gastropod Urosalpinx cinerea (SayY) (Carriker 1978), and
cephalopod Octopus vulgaris Cuvier (Nixon et al. 1980).
A secretion weakens the shell surface by initially solubilizing
the nonmineralized intercrystalline organic matrix between
individual mineral cores of shell units, then dissolves exposed
mineral cores; dissolution of organic matrix and mineral
cores then proceeds at more or less equal rates, solubiliza-
tion of the organic matrix ahead of mineral cores, the latter
frequently irregular and pitted. The secretion of the accessory
boring organ of U. cinerea, hypothesized to contain a com-
bination possibly of HC1, chelating agent, and enzyme
(Carriker 1981) could produce the differential dissolution
observed ultrastructurally. Similarity of the pattern of
etching produced in shell penetration of P. websteri,
T. lampas, U. cinerea, and O. vulgaris suggests the existence
of a generically similar chemical mechanism in the shell-
penetrating Eumatozoa.

COMPOSTING OF BLUE CRAB SCRAP:
PROBLEMS AND SOLUTIONS

THOMAS P. CATHCART, FRED W.
WHEATON AND RUSSELL B.
BRINSFIELD

Department of Agricultural Engineering
University of Maryland
College Park. Maryland 20 742

Disposal of solid waste from blue crab processing plants
became a major problem in Maryland with the closing of
dehydrating plants. The dehydrated crab waste (scrap) was
ground and sold for chicken feed. Presently, the scrap is
disposed of in landfills; however, risk of ground water
pollution and operational problems of placing crab scrap
in landfills limits landfilling to a temporary solution. Com-
posting of the crab scrap is a possible method of stablizing
the waste and producing a useful soil additive for farmers,
gardeners, the potted-plant industry, and others. Composting
of crab scrap requires special provisions to eliminate noxious
odors and prevent nuisance problems from developing.
Studies to date have shown that the crab scrap pH must be
maintained below 7.5 during composting, aeration must be
supplied during part of the composting cycle, and a source
of additional carbon must be added to the scrap. Solutions
to these problems and methods of composting have been
developed which produce high quality compost without
noxious odor production.

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

OPTIMUM SALINITY REGIME FOR OYSTER

PRODUCTION ON LOUISIANA'S

STATE SEED GROUNDS

MARK CHATRY AND R. J. DUGAS

Lyle S. St. Amant Marine Laboratory
Grand Terre Island. Louisiana 70358

Increased salinities have drastically reduced the produc-
tive portion of Louisiana's public oyster seed grounds.
Controlled freshwater diversions from the Mississippi River
have been utilized or are now being planned in an attempt to
reduce salinities and thereby reestablish formerly productive
reefs. These diversions offer an unprecedented opportunity
to manipulate salinities over a vast estuarine area for maxi-
mizing seed oyster production. The purpose of this study
was to determine the optimum annual salinity regime, using
historical data, for the production of seed oysters on
Louisiana's seed grounds.

Salinity, spatfall, and seed oyster production data from
three stations on Louisiana's productive seed grounds,
1971 — 1981, are presented. Salinity in the setting year was
the prime factor determining production of seed oysters.
Both high and low salinity extremes resulted in poor seed
production. Insufficient setting was blamed for poor pro-
duction at the low salinities and it was speculated that
numerous organisms associated with the high salinities
caused heavy mortalities in recently set oysters. The optimum
annual salinity regime was derived from all of the year/
station salinity regimes which were followed in the ensuing
year by good seed oyster production. This optimum regime
accounts for the salinity dependent factors which limit seed
production.

GENE STRUCTURES OF ATLANTIC COAST POPULATIONS
OF THE BLUE CRAB CALLINECTES SAPIDUS RATHBUN

TIMOTHY J. COLE

University of Maryland Center for
Environmental and Estuarine Studies
Horn Point Environmental Laboratories
Box 775. Cambridge. Maryland 21613

Recent research has indicated that larvae of blue crabs
are probably flushed from their parent estuary. Develop-
ment continues in offshore waters, after which late-stage
larvae or post-larvae return to the estuaries. A genetic study
of blue crab populations was undertaken to determine if
there is sufficient gene exchange among estuaries to prevent
differentiation. Horizontal starch-gel techniques were used.
Statistical analyses of frequencies of polymorphic loci indi-
cate that blue crab populations south of Cape Hatteras are
more genetically similar to each other than to those north
of that cape.

NATICID SNAIL PREDATION IN NEW ENGLAND: THE

EFFECTS OF LUNATIA HEROS ON THE POPULATION

DYNAMICS OF MY A ARENARIA AND

MACOMA BALTHICA

JOHN A. COMMITO

Department of Biology

Hood College

Frederick, Maryland 21 701

The naticid snail predator Lunatia heros (Say) and two
of its bivalve prey species, Mya arenaria Linne and Macoma
balthica (Linne), were studied at an intertidal site in eastern
Maine. The M. arenaria population was comprised largely of
newly recruited individuals. Survivorship was low (3.5%/y)
until the sixth year and increased thereafter. Lunatia heros
preyed upon only those individuals of M. arenaria < 30 mm
long. At that length the bivalve reached a size or depth
refuge from predation. It delayed reproduction until it was
4 years old (20 mm long) and allocated its resources to rapid
early growth instead (4.9 mm/y for the first 5 y).

The dynamics of the population of M. balthica were
different. There was a larger proportion of older individuals
of M. balthica, and survivorship was higher (76.3%/y for the
first 5 y). Macoma balthica grew to a length of 25 mm and
never reached a size refuge. All sizes were susceptible to
attack by L. heros, but the deeper burrow of M. balthica
relative to individuals of M. arenaria of the same size may
have afforded it some protection from predation. Macoma
balthica grew slowly (2.7 mm/y for the first 5 y) and
diverted its resources into reproduction at a younger age
(3 y) and smaller size (10 mm). These different life-history
patterns and the possible relationship between bivalve
resource allocation and refuges from predation are discussed.

THE EFFECTS OF POLLUTANTS ON LARVAL DEVELOPMENT
OF THE BLUE CRAB CALLINECTES SAPIDUS RATHBUN

J. D. COSTLOW AND C. G. BOOKHOUT

Duke University Marine Laboratory
Beaufort, North Carolina 28516

Since our initial rearing of all larval stages of the blue crab
Callinectes sapidus from hatching to the juvenile crab, we
have investigated the way in which a variety of pollutants
may affect the survival, duration, and frequency of abnor-
mality of larvae of this important commercial species.
Having established the optimum temperatures and salinities
required for total development, we have investigated the
way in which a number of commonly used pesticides and
heavy metals affect development, either singly or in combina-
tion with those temperatures and salinities which are known
to impose a stress on the developing larvae. Included among

Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

the pesticides have been studies on Malathion, Methoxychlor.
Mirex, Kepone, and Dimilin. Studies on the effects of heavy
metals have included cadmium and mercury.

Summary data involving these studies are presented and
discussed. In all cases, small amounts of each of the chemicals
tested reduced survival of the larvae. Even at "sublethal"
levels, abnormalities in development were observed. In
general, the larval stages were far more sensitive to pollutants
than were the juvenile or adult crabs and any consideration
of "water quality" should take into consideration this
essential portion of the life cycle of the blue crab and the
sensitivity of the various larval stages to extremely minute
amounts of pollutants.

ANALYSIS OF LOCAL POPULATIONS OF THE BLUE CRAB
CALLINECTES SAPIDUS RATHBUN

L. EUGENE CRONIN

Chesapeake Research Consortium

4800 Atwell Road

Shady Side, Maryland 20764

The catch of blue crabs and composition of that catch
fluctuate rapidly and widely over time. Useful estimation of
local availability, size structure, and sex composition is,
however, essential for understanding and for management
of the species. A procedure of obtaining such information
is described and discussed. It involves detailed catch infor-
mation from the best of samplers (selected professional
crabbers) accompanied by appropriate quantitative observa-
tion at frequent intervals on the composition of the catch.
These can provide useful estimates of the number of each
class of crab available per man day throughout the crabbing
season. The advantage and limitations are considered.

CHEMORECEPTION AND LIFE HISTORY OF
STYLOCHUS ELLIPTICUS (GIRARD)

PETER DANIEL 1 , TIMOTHY COLE 1 ,
AND DANIEL RITTSCHOF 2

1 Horn Point Environmental Labs
University of Maryland
Cambridge, Maryland 21613, and
"College of Marine Studies
University of Delaware
Lewes, Delaware 1 9958

Stylochus ellipticus, a flatworm indigenous to the
Atlantic coast of the United States, preys on oyster spat and
barnacles. Adults have almost inflexible prey preferences.
Little is known about early life stages. A prey chemolocation
hypothesis was tested to explain ability of S. ellipticus to
locate and discriminate prey species. Also, these studies
initiated examination of life history and distribution of
5. ellipticus in Chesapeake Bay.

Three apparatuses (chemossayer, Y-maze, and choice-
chambers) were used to test adults for chemoreception.
Effects of various environmental and biotic factors onchemo-
reception were tested. The Atlantic oyster drill Urosalpinx

cinerea (Say), an ecological analogue with an extensively
studied chemobiology, was used to verify apparatus effec-
tiveness and stimulus and control water attractiveness.
Survivorship of larvae in nutrition and substrate preference
settlement studies was determined. Distribution of S.
ellipticus in Chesapeake Bay was determined from oyster
bar survey reports (1980—81), occurrence in oyster hatch-
eries (1980—81), and prior fouling plate studies (1963-65)
(Shaw 1967).

Studies of U. cinerea verified effectiveness of apparatuses
and of stimulus and control water. Chemoreceptive behavior
was indicated only in choice-chamber studies as long
response time of adults rendered other apparatuses ineffec-
tive. Light and starvation modified prey search. Stylochus
ellipticus has a Gotte's larva which appears to be non-
feeding and metamorphoses only on prey substrates. Though
flatworm and prey densities often correlate, there were
several instances of uninfested prey populations.

Adults of S. ellipticus appear to prioritize behavior: (1)
reproduction vs. prey search, and (2) prey search vs. escape.
Barriers to larval dispersal probably allow some prey popula-
tions to escape infestation. Earlier, nonreproductive life
stages may influence prey preference establishment.

EFFECT OF CRAB POT WIRE TREATMENT ON CRAB POT
FOULING IN CHESAPEAKE BAY, MARYLAND

RAY C. DINTAMAN AND J.F. CASEY

Tidewater Administration, Maryland
Department of Natural Resources
Annapolis, Maryland 21401

It has been presumed that fouling on crab pots reduces
the catch rate and contributes to a shortened fishing life or
premature loss of the pot. Groups of standard anode pots,
standard anode pots painted with an anti-fouling paint, and
vinyl pots were compared for rate of fouling and catch. Crab
pots treated with the anti-fouling paint fouled the least.
Their fouling rate was 83% less than vinyl pots and 75% less
than standard anode pots. Pots treated with anti-fouling
paint accounted for 42% of the total crabs caught during
the study. This study suggests that treatment of standard
anode pots with anti-fouling paint could increase not only
catch, but also pot life.

AN OYSTER CULTCH COMPARISON :
CLAMSHELL VS. LIMESTONE

CHARLES N. DUGAS AND M. CHATRY

Lyle S. St. Amant Marine Laboratory
Grand Terre Island, Louisiana 70358

On 15 April 1981 four 70- X 70-cm trays containing equal

National Shellfisheries Association, Baltimore, Maryland

Abstracts. 1982 Annual Meeting, June 14-17, 1982

volumes of clamshell and graded crushed limestone were
placed on the bottom at each of 10 stations in the Barataria
Bay system of southeast Louisiana. At the end of 3 months
two trays and their contents from each station were retrieved
and replaced with two trays containing fresh material. After
the following three months all trays were retrieved. Thus,
the cultch materials were exposed to spat set for two suc-
cessive 3-month periods and for one 6-month period. Spat
set (spat/liter of cultch) was determined by counting live
and dead spat on each piece of cultch material. The overall
mean spat set/liter was 57.9 for limestone and 25.1 for
clamshell. This ratio of approximately 2:1 also held true
when the data were analyzed for each time period. Relative
survival was slightly higher on clamshell; however, because
of the greater set on limestone, there was still approximately
twice the number of live spat on limestone as on clamshell.
At current prices crushed limestone is approximately 60%
higher than clamshell; however, since spat set on limestone
was greater, the cost, using average prices, was about
$0 .50/ 1 ,000 spat on limestone and $0 .70/ 1 ,000 onclamshell.

INCIDENCE OF PATHOGENIC BACTERIA IN THE

BLUE CRAB CALLINECTES SAPIDUS RATHBUN

AND THE AMERICAN OYSTER CRASSOSTREA

VIRGINICA (GMELIN)

ELISA L. ELLIOT AND
RITA R. COLWELL

Department of Microbiology
University of Maryland
College Park, Maryland 20742

Blue crabs (Callinectes sapidus) and American oysters
(Crassostrea virginica) were analyzed for the presence of
human pathogenic bacteria. Live and cooked crabs, freshly
picked crabmeat, and live, shucked, and washed oysters
were obtained from a Maryland processing plant in the
winter and spring of 1 981 -82. Cans of pasteurized crabmeat,
purchased in Washington, DC, area stores, were also included
in the study. All samples were subjected to standard plate-
count determination and enrichment for the detection of
specific pathogens. Sample analyses revealed low numbers
of Staphylococcus aureus Rosenbach, Vibrio parahaemoly-
ticus (Fujino et al.), other halophilic Vibrio spp., Aeromonas
hydrophila (Chester), fecal coliforms, and presumptive
Clostridium perfringens (Veillon and Zuber) spores; Vibrio
cholerae Pacini and Salmonella spp. were not detected.
Excluding S. aureus, all of the pathogens were present in
highest numbers in the live crabs and oysters, suggesting
that processing is effective in controlling the numbers of
pathogens present in these foods.

PREDATION ON SPAT OF THE AMERICAN OYSTER

CRASSOSTREA VIRGINICA (GMELIN) BY THE

AMERICAN LOBSTER HOMARUS AMERICANUS

MILNE -EDWARDS. THE ROCK CRAB CANCER

IRRORATUS (SAY), AND THE MUD CRAB

NEOPANOPE SA YI (SMITH)

R. W. ELNER 1 AND R. E. LAVOIE 2

Department of Fisheries and Oceans

Biological Station

St. Andrews, New Brunswick

Canada E0G 2X0, and

Department of Fisheries and Oceans

Fisheries Research Branch

Halifax, Nova Scotia, Canada B3J 2S7

Predation by lobsters, rock crabs, and mud crabs on
oyster spat was compared in the laboratory at 13°C. Rock
crabs (32- to 107-mm carapace width, CW) preyed on
oysters up to 30 mm length, although they preferred smaller
oysters. Preferred prey size increased with rock crab size.
Lobsters (55- to 98-mm carapace length) demonstrated a
broad preference for oysters of 1 0- to 25-mm length. Oysters
up to 35-mm length were vulnerable to the lobsters. Preda-
tion rate was highly variable but generally increased with
predator size. Maximum mean lobster and rock crab preda-
tion rates were 4.5 and 28.0 oysters/predator/day, respec-
tively. Mud crabs (14- to 23-mm CW) and rock crabs (32- to
58-mm CW) feeding on oysters (2- to 9-mm length) attached
to spat collectors ate approximately 0.5 oyster/predator/day.

Lobsters used their mouthparts or chelae to open oysters
by indiscriminate crushing. Rock crabs generally crushed
the umbo, chipped away the shell margin, or punctured the
prey shell. Mud crabs and rock crabs opened oysters still
attached to the spat collector. Oyster fragments were found
in the stomachs of 88 (44%) of 201 rock crabs collected
around oyster beds in Caraquet Bay, New Brunswick.

SEASONAL OCCURRENCE OF THE LARVAE OF

CALLINECTES SAPIDUS RATHBUN IN

DELAWARE BAY

CHARLES E. EPIFANIO. C. C.
VALENTI AND A. E. PEMBROKE

College of Marine Studies
University of Delaware
Lewes, Delaware 19958

Blue crab larvae were collected weekly at a station in the
mouth of Delaware Bay over a 16-wk period beginning in
late June 1979. Collections were made with a 0.3-m Clark-
Bumpus Sampler; discrete samples were taken at the surface,
at 12 m, and at the bottom (25 m). On each sampling date,
larvae were collected at the three depths every 3 h over one

90 Abstracts, 1982 Annual Meeting, June 14-17. 1982

National Shellfisheries Association, Baltimore, Maryland

tidal cycle. Only Stage I zoeae and megalopae were collected
during the course of the investigation. Peak abundance of
Stage I occurred during late July and early August while
peak occurrence of megalopae was observed 5 wk later.
Stage I larvae were most abundant in seaward-flowing sur-
face water and megalopae were distributed throughout the
water column. We concluded that blue crab larvae are
exported from the Bay as Stage I zoeae, undergo subsequent
zoeal development on the continental shelf, and return to
the estuary as megalopae.

Adult of Crassostrea virginica were collected from 51
sites in Chesapeake Bay and its tributaries. Samples were
analyzed for heavy metal, polychlorinated biphenyl (PCB),
and pesticide contamination. Ranges, medians, means, and
standard deviations were determined for the Maryland por-
tion of Chesapeake Bay and for some major river systems.
Trends indicated by the 1980 data are discussed. Data are
compared to previously collected data.

CHARACTERISTICS OF FECAL RIBBONS FROM JUVENILES

OF CRASSOSTREA VIRGINICA (GMELIN) FED

PHAEODACTYLUM TRICORNUTUM BOHLIN

WITH AND WITHOUT THE ADDITION OF

SILT: PRELIMINARY OBSERVATIONS

JOHN W. EWART AND
MELBOURNE R. CARRIKER

College of Marine Studies
University of Delaware
Lewes, Delaware 19958

Two size classes of Crassostrea virginica were (edPhaeo-
dactylum tricornutum at two cell concentrations with and
without the addition of silt. The experimental treatments
included 3-g and 21-g oysters, algal concentrations of 1.0
X 10 4 cells/ml and 1 .0 X 10 s cells/ml, and either natural or
oxidized Broadkill River silt at a concentration of 50 mg/C.
Each treatment was tested in replicate feeding trials lasting
24 h. Microscopic examination of fecal ribbon contents
from oysters fed at the low algal concentration showed that
the addition of silt resulted in a marked reduction in the
number of whole cells of P. tricornutum. At the higher algal
concentration the addition of silt had no effect on reducing
the number of whole cells in the fecal ribbons. No differ-
ences in the effect were found between oyster size classes.
SEM examination of all fecal material indicated that silt-
treated samples were different in appearance and composi-
tion from those fed algae alone. The implications of silt
additions in improving the nutritive value off. tricornutum
are discussed.

HEAVY METAL, POLYCHLORINATED BIPHENYL, AND

PESTICIDE LEVELS IN CRASSOSTREA VIRGINICA

(GMELIN) FROM CHESAPEAKE BAY

MARY JO GARREIS AND
F. A. PITTMAN

Office of Environmental Programs
Department of Health and Mental
Hygiene, 201 W. Preston Street
Baltimore, Maryland 21201

REDUCTION OF DISSOLVED ORGANICS IN BLUE CRAB
PROCESSING PLANT EFFLUENT

EUGENE L. GEIGER, RUSSELL B.
BRINSFIELD AND FRED W. WHEATON

Department of Agricultural Engineering
University of Maryland
College Park. Maryland 20742

Blue crab processing plants have difficulty meeting dis-
charge guidelines for federal and Maryland state liquid
effluents. Conventional treatment systems (e.g., foam flota-
tion or aerated lagoons) do not represent viable options
because of severe land and cost constraints. Research was
initiated to develop: 1 ) a cost effective effluent treatment
system and 2) a system producing effluent of sufficient
quality to meet discharge guidelines. An attempt was made
to utilize ultraviolet light as a substitute for chlorination.
Crab cooking retort water, diluted to a 5% strength, was
used as a consistent feed solution containing a high level of
dissolved organics. Chemical floculation (with aluminum
sulfate, ferric chloride, or ferrous sulfate), foam fractiona-
tion, and aerobic biological treatment were examined in the
laboratory using this solution to determine the most promis-
ing treatment method. Because of the high dissolved organics
concentration in the effluent, aerobic biological treatment
proved to be the most effective treatment method. Various
retention times in a sequential biological reactor were
studied. A significant reduction in dissolved organic concen-
trations was achieved, but substantial concentrations of col-
loidal particulates were produced. Filtration with a fine sand
filter greatly reduced the particulate concentrations. Final
polishing by activated carbon absorption produced effluent
transmission values in the range necessary for effective dis-
infection by ultraviolet light. Water quality parameters were
monitored between each treatment step. The quality of the
water leaving the scale model system met federal and Mary-
land state discharge limitations.

National Shellfisheries Association, Baltimore, Maryland

Abstracts. 1982 Annual Meeting, June 14-17, 1982 91

MORPHOMETRY PATTERNS IN INTERT1DAL BIVALVES

REGINALD B. GILLMOR AND
HERBERT HIDU

Ira C. Darling Center
Walpole, Maine 045 73

For several families of intertidal gastropods Vermeij
(1973) has demonstrated low-to-high shore gradients in shell
morphology which he interpreted in terms of adaptive
responses to the dominant physical stresses of the shore
environment. Evidence from a variety of studies suggests
that similar responses may occur in bivalves. The present
study examined this question further. Juveniles of six bivalve
species (Argopecten irradians [Lamarck] , Modiolus modiolus
[Linne] , Ostrea edulis Linne, Mytilus edulis Linne, Crassos-
trea virginica [Gmelin] , and Geukensia demissa [Dillwyn] )
were grown at various tidal levels on a natural shore and in
a laboratory tidal simulator. At the end of the treatment
period, the bivalves were sacrificed and each specimen was
measured for maximum shell dimension (MSD: length in
the mussels, height in the other species) and width; dry meat
and dry shell weights were also determined. Three morpho-
metric ratios were calculated and compared among species
and treatment groups: shell weight/(MSD X width) as an
index of relative shell thickness; MSD/width as an index of
relative shell globosity; and meat weight/shell weight. Bivalves
that were grown intertidally tended to have thicker and
more globose shells. These tendencies did not necessarily
correlate with naturally occurring or experimental intertidal
levels. Intertidal meat/shell ratios, however, corresponded
closely to natural shore position; the lower-shore species
had the lowest ratios and the higher-shore species had the
highest. We concluded that inter-specific and, in some cases,
intra-specific low-to-high shore gradients in morphometric
relationships are present in bivalves.

NONPLANKTOTROPHIC LARVAL DEVELOPMENT OF
TWO SPECIES OF CONTINENTAL SHELF BIVALVES

M. CASTAGNA' AND J. KRAEUTER"

x Dept. of Oyster Culture, NJAES
Cook College, Rutgers University
New Brunswick, New Jersey 08903
VIMS. Wachapreague, Virginia 23480

Larvae of Periploma leanum (Conrad) and Astarte cas-
tanea (Say) were reared under laboratory conditions. The
larval stages of both species are lecithotrophic and have low
dispersal capabilities. Spawning was induced in P. leanum
with thermal stimulation and the addition of a gamete sus-
pension following a period of intensive feeding. Individual
eggs (dia. = 130 jim) were released inside of two-layered
capsules. The outer gelatinous layer rapidly expanded and,
within 24 hours, completely dissipated. After 4 to 6 days,

straight-hinge larvae emerged from an opening at the
restricted end of the inner capsule. After a planktic stage of
< 24 h, the larvae (length = 170 p.m) assumed an inactive
benthic existence; a functional foot was not observed until
15 to 18 days after fertilization. At no time during larval or
early postlarval development were byssal threads observed.
Astarte castanea was induced to spawn with thermal stimu-
lation and the addition of a gamete suspension. Individual
eggs (dia. = 170 yum) were released inside of double-walled,
adhesive capsules. Prodissoconch I formation was extremely
slow. The first sign of valve formation was observed after 6
to 10 days while the larvae rotated within the capsules.
Movement within the capsule ceased between 8 and 15 days
after fertilization when the valves first completely enclosed
the soft tissues and closed against one another along their
free margins. Between 22 and 26 days, young of A. castanea
broke out of their capsules by pushing forcefully with their
foot against the inner wall of the capsule. They emerged as
benthic juveniles (In. = 240 (im). As a result of the adhesive
nature of the encapsulated stages, the larval dispersal capa-
bility of this species is estimated to be on the order of a few
centimeters.

THE ROLE OF THE VENTRAL PEDAL GLAND IN

FORMATION OF AN EGG CAPSULE BY THE

MURICID GASTROPOD EUPLEURA CAUDATA

ETTERAE B. B. BAKER 195 1: AN INTEGRATED

BEHAVIORAL, MORPHOLOGICAL, AND

HISTOCHEMICAL STUDY

GREGORY L. GRUBER

College of Marine Studies
University of Delaware
Lewes, Delaware 19958

Several researchers described formation of egg capsules
by females of a few neogastropods, but this process is still
not well understood. Spawning behavior of females defined
discrete times to sample egg capsules and spawning females
before ventral pedal gland activity (VPGA), after peristaltic
molding during VPGA, and after VPGA. Structure of these
egg capsules and ventral pedal glands of females was examined
with dissections, histology, polarizing microscopy, and
histochemistry. Egg capsules before VPGA were ovoid,

soft, and flexible. After peristaltic molding during VPGA,
egg capsules were roughly shaped, loosely attached to
a hard substratum, and still soft and flexible. Egg capsules
after VPGA were completely shaped, firmly attached to a
hard substratum, but now hardened and resilient. The apical
plug, embryo chamber, and multilyatered fibrous wall
of egg capsules before, during and after VPGA had similar

92 Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

morphologies. Histochemical composition of the wall of egg
capsules before VPGA and after peristaltic molding during
VPGA differed from that of the wall of the egg capsules
after VPGA. The wall of whole egg capsules that were sam-
pled before VPGA and exposed to filtered seawater for 5
days were soft, flexible, and showed no histochemical
changes. These observations suggested that the ventral pedal
gland molded an egg capsule into its final species-specific
shape, firmly attached it to a hard substratum, chemically
hardened the wall of the egg capsule, but did not secrete
any layers of its wall. The ventral pedal gland has a columnar
epithelium, two types of epithelial goblet cells, clusters of
subepithelial gland cells, and a thin layer of circular and
longitudinal muscle fibers between the epithelium and these
gland cells. Each goblet cell type secreted different sulfated,
acid mucosubstances that may act as lubricants during mold-
ing of egg capsules. Subepithelial gland cells may secrete a
noncarbohydrate, nonprotein substance that hardens the
wall of the egg capsule.

SOME RELATIONSHIPS AFFECTING GROWTH OF SEED

OF THE HARD CLAM MERCENARIA MERCENAR1A

IN RACEWAYS

NANCY H. HADLEY 1 AND JOHN J.
MANZI 2

Grice Marine Biological Laboratory
216 Ft. Johnson Rd.
Charleston, South Carolina 29412
Marine Resources Research Institute
Charleston, South Carolina 29412

Seed clams (y size = 3.9 mm) were maintained in race-
ways for 6 months at densities corresponding to 740, 2220,
6660, and 19980 clams/m 2 . Each density was replicated
eight times in the raceways and the highest and lowest densi-
ties were replicated four times in subtidal field controls.
Raceway clam populations were stocked in four different
positions relative to water flow and in 19 different positions
relative to total raceway biomass. Although nominal flow
rate was constant, effective flow rate (water volume/clam
volume/minute) was different for each replicate and decreased
as clam biomass increased. Temperature and salinity were
measured daily and inflow and outflow chlorophyll-a were
monitored monthly from February to August 1981 to deter-
mine growth and survival. Single classification ANOVA fol-
lowed by SNK tests between means showed that growth
was significantly reduced at the highest density in both the
raceway and the field. The lowest density exhibited greater
growth in the raceway than in the field, while the highest
density showed no difference in growth between the two
locations. In the raceway, growth rate was inversely propor-
tional to distance from water inflow and to effective density

(# clams/unit water). Although clams at the highest density
consistently removed the greatest amount of chlorophyll-a,
less chlorophyll was removed per clam as density increased.
Growth was highly correlated with stripping rate (/ig
chlorophyll-a/clam/day) and with effective water flow rate.
These relationships are discussed and some implications for
management of raceways in mariculture systems are made.

MLXED-FUNCTION-OXYGENASE ENZYME SYSTEMS:
PURPOSE AND POSSIBLE DELETERIOUS INTER-
ACTIONS WITH ORGANIC POLLUTANTS
IN THE BLUE CRAB

ROBERT C. HALE

Virginia Institute of Marine Science
The College of William and Mary
Gloucester Point, Virginia 23062

Mixed-function-oxygenases (MFO) are enzyme systems
which have evolved in organisms to enable them to eliminate
foreign compounds taken in from their environment. Often
these compounds are toxic and lipophilic, possessing high
accumulative potential (e.g., polynuclear aromatics, poly-
chlorinated biphenyls, and chlorinated organic pesticides);
therefore, they must be metabolized to biologically inactive,
excretable forms. Occasionally, however, the resulting
metabolites formed by the MFO system are more harmful
than the parent compounds; some are potent carcinogens.
Recent work has shown that the activity of the MFO sys-
tem is greatest in mammals and decreases in fish, crustaceans,
and mollusks, in that order. The enzyme system is also respon-
sible for the synthesis and breakdown of certain steroid
hormones. The molting hormone in crustaceans is believed
to be a steroid compound. The activity of MFO in female
blue crabs has been shown by others to be inversely related
to the levels of crustecdysone, when examined over the
course of a molt cycle. Elevated levels of aromatic hydro-
carbons, caused by greater utilization of coal reserves and
increased industrialization, are of concern to scientists.
These and other pollutants have been found by workers to
induce higher levels of MFO activity, and also to inhibit
molting and limb regeneration in crabs. Levels of toxic
organic compounds in the blue crab population of lower
Chesapeake Bay are being determined using glass capillary
gas chromatography and mass spectrometry. Differential
abilities to metabolize aromatic compounds that may exist
between molt and sex groups will be examined.

ESTIMATES OF JUVENILE BLUE CRAB
ABUNDANCE IN TEXAS BAYS

PAUL C. HAMMERSCHMIDT

Texas Parks and Wildlife Department
Rt. 1, Box 368, Seadrift, Texas 77983

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

Blue crab populations were monitored November 1 977—
December 1981 by Texas Parks and Wildlife Department
personnel using 18-m bag seines in the Galveston, Matagorda,
San Antonio, Aransas, Corpus Christi, upper and lower
Laguna Madre Bay systems. Seine samples and hydrological
data were taken monthly at randomly selected stations in
each of the sampled bay systems. Catch-per-unit-of-effort
(CPUE), calculated as number of crabs/ha, as well as water
temperature and salinity values are presented. These data
were examined utilizing a 2-way ANOVA. Similarities in
CPUE, water temperature, and salinity were examined
between years and seasons within bay systems.

THE SURF CLAM ALONG THE NEW JERSEY COAST:
POPULATION SIZE. RECRUITMENT, GROWTH RATES

HAROLD H. HASKIN, ERIC S.
WAGNER AND MITCHELL L.
TARNOWSKI

Department of Oyster Culture

N.J. Agricultural Experiment Station

Rutgers the State University

New Brunswick, New Jersey 08903

Over the last 10 years there has been regular and general
settling of surf clam larvae along the New Jersey coast but.
as indicated in earlier reports, mortality rates in early juve-
niles are high and survival beyond the first summer is com-
paratively rare. Exceptions to this will be discussed with
emphasis on the 1976 year class which approximately
doubled the standing stock in New Jersey waters. Since
major portions of this year-class survived in areas where
earlier year-classes were wiped out by anoxic waters in
1976, we have a unique opportunity to determine the effects
of a variety of environmental conditions on growth rate.
Results of some of these determinations will be presented,
as will the most recent stock assessment.

GROWTH PERFORMANCE OF CYTOCHALAZIN-INDUCED

TRIPLOIDS OF AMERICAN OYSTERS AND

SOFT-SHELL CLAMS

HERBERT HIDU, STANDISH ALLEN
AND JON STANLEY

Department of Zoology
University of Maine at Orono
Orono, Maine 04469

We conducted extensive laboratory and field performance
experiments in 1982 with 3-yr-old triploids of the American
oyster Crassostrea virginica (Gmelin) and yearlings of the
soft-shell clam Mya arenaria Linne. The Crassostrea triploids,
which were created at meiosis I, grew significantly faster
than the diploid controls, whereas those created later in the
meiotic cycle exhibited no growth advantage over the dip-

loids. The Mya triploids exhibited no growth advantage over
diploid controls. Triploidy did not block gametogenesis in
either species. Optimal methods are discussed for determin-
ing the consequences of polyploidy in marine bivalves.

PREDATION BY BLUE CRABS AND SPOT ON INFAUNAL
COMMUNITIES IN CENTRAL CHESAPEAKE BAY

ANSON H. HINES AND KATHRYN L.
COM TO IS

Chesapeake Bay Center. Smithsonian
Institution. P.O. Box 28
Edgewater, Maryland 21037

The impacts of predation by blue crabs (Callinectes
sapidus Rathbun)and spot(Leiostomusxanrhuri(sLacephde)
on infaunal communities were compared for mud and sand
sediments in the Rhode River, a typical subestuary of central
Chesapeake Bay. The two species are the dominant benthic
predators in the system, and their foraging activities from
June to October correlated with the sharp seasonal decline
in infaunal density and standing crop. Analysis of stomach
contents showed that crabs preyed primarily on whole
clams, whereas spot fed mainly on clam siphons and several
species of polychaetes. Turnover rates of infaunal prey were
estimated based on the density of predators taken in otter
trawls, the weight of their stomach contents, and the weight
of the standing crop of infauna. For total infauna, turnover
rates were low (1— 7%/month) early in the season, when the
standing crop was high; but turnover was high (30-60%/mo)
in the top 5 cm of sediment late in the season, when the
standing crop was low. For small clams, polychaetes, and
amphipods in the top 5 cm of sediment, predation pressure
by crabs and spot accounted for extremely high turnover
rates (more than 100%/mo), whereas larger, deep-burrowing
clams had turnover rates < 3%/mo. Experiments using pre-
dator exclusion cages resulted in significantly higher densities
of total infauna, clams, and some species of polychaetes
within the cages than outside the cages. Survival of out-
planted clams (Macoma balthica [Linne] ) was significantly
higher in buckets with predator exclusion cages than in
buckets without predator exclusion cages. Predation by
blue crabs appears to have a major impact on small, surface-
dwelling clams, whereas spot predation has a more general
impact on clam siphons and a variety of invertebrates living
in the surface sediment. Turnover of infauna in the surface
sediment is very rapid.

94 Abstracts. 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

OCEANOGRAPHY OF THE SOUTHEASTERN BERING SEA

AND RECRUITMENT PROCESSES IN TWO SPECIES

OF TANNER CRAB

LEWIS S. INCZE

School of Fisheries WH-10

College of Ocean and Fishery Sciences

University of Washington,

Seattle, Washington 98195

Potential factors affecting the distribution and survival
of the pelagic larvae of two species of tanner crabs, Chiono-
ecetes bairdi Rathbun and C. opilio (Fabricius), that inhabit
the wide continental shelf of the eastern Bering Sea were
investigated as part of a large multi-institutional oceano-
graphic program. The objective was to evaluate the relative
importance of pelagic events in determining spatial patterns
of recruitment to the benthos. The study emphasized the
description of cause-and-effect relationships between physi-
cal processes (mixing and transport) and biological (plank-
tonic) conditions which affect feeding success and the
ultimate survival and distribution of the larvae. Information
on the timing of hatch-out, rates of growth and development,
feeding physiology, and inter-annual differences in patterns
of spatial distribution and relative abundance of the larvae
are provided. How these data relate to regional oceanographic
processes and their potential impact on population distribu-
tion and age structure are stressed.

SPECIES-SPECIFIC DIFFERENCES IN THE

MEGALOPAL DISTRIBUTIONS RELATED

TO WATER DENSITY PARAMETERS

DAVID F. JOHNSON

Department of Oceanography
Old Dominion University
Norfolk. Virginia 23508

The megalopae of 10 brachyuran crabs were sampled
from July through September 1980 in the lower Chesapeake
Bay and adjacent coastal waters. The megalopae are assigned
to three apparent groups: retained estuarine. expelled estu-
arine, and retained coastal recruitment types. The megalopae
of estuarine species such as Hexapanopeiis angastifrons
( Benedict and Rathbun), Neopanope sayi (Smith ). Panopeus
herbstii H. Milne-Edwards, and Pinnotheres ostreum Say are
retained in estuarine epibenthic waters. The larvae of some
estuarine species such as Callinectes sapidus Rathbun, Uca
spp.. and Pinnixa spp. are expelled from the estuary, resul-
ting in maximum megalopal concentrations on the shelf. Of
the retained coastal species. Portunus spp. and Cancer irrora-
tus Say are not abundant in the neuston of shelf waters,
while Libinia spp. are most abundant in the epibenthos of
near-shelf waters. The megalopae of 4 species show signifi-
cantly different vertical distributions between stratified and

homogenous water columns. Megalopae were not found to
aggregate within pycnoclines.

MECHANISM OF SHELL PENETRATION BY THE

BURROWING BARNACLE TRYPETESA LAMP AS

(HANCOCK), (CIRRIPEDIA: ACROTHORACICA):

AN ULTRASTRUCTURAL STUDY

TODD C. KAMENS

College of Marine Studies
University of Delaware
Lewes. Delaware 19958

Trypetesa lampas is a soft -bodied, free-living cirriped that
burrows in empty shells of gastropods inhabited by hermit
crabs. Portions of this burrow are commonly lined with a
limy, white material. Individuals of T. lampas were obtained
from shells of Lunatia heros (Say) and Polinices duplicatus
(Say) collected in the vicinity of Woods Hole, Massachusetts.
Specimens of the mantle surface and burrow wall were
examined with scanning electron microscopy to determine
the mechanisms of shell removal and lining formation within
the burrow by T. lampas and to correlate these activities
with the microanatomy of the external mantle surface of
the barnacle. Results confirm earlier hypotheses that bur-
rowing by T. lampas is achieved through a combination of
chemical and physical processes. Ultrastructural examination
of fractures through the burrow reveal a gradual, shell-
weakening process in which prismatic material within the
surrounding gastropod shell is softened by preferential dis-
solution of inter- and intra-crystalline matrix and subsequent
solubilization of the bare calcareous prisms. Examination of
thin sections through the mantle cuticle disclosed minute
pore canals through which shell-dissolving secretions of the
barnacle could be released. Dissolution of shell by T. lampas
appears to be linked to the molt cycle, with most extensive
stages of dissolution being observed in burrows of specimens
that have just molted. Soft material remaining on the wall
of the burrow after molting is removed with sharp spines
covering the external surface of the barnacle's mantle. This
material is subsequently used by T. lampas to thicken exist-
ing parts of the lining and add new linings in areas that no
longer fit snugly.

TRACE METALS IN SHELLFISH AND
GROWING AREA DESIGNATION

JEFFREY KASSNER

Department of Environmental Protection
Town of Brookhaven
Patchogiie, New York 11772

The level of coliform bacteria, as set forth by the National
Shellfish Sanitation Program (NSSP), is the water quality

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982 95

standard used to classify shellfish growing areas. It is the
standard by which shellfish harvesting is regulated. Port
Jefferson Harbor, NY, a moderately industrialized embay-
ment of Long Island Sound, and Setauket Harbor, a more
urbanized tributary basin of Port Jefferson Harbor, both
have areas classified as certified (shellfishing permitted) and
as uncertified (shellfish prohibited). Sediment analyses of
the two harbors suggest that noncoliform pollutants, particu-
larly trace metals, are present. Because of public health
concerns, the hard clam Mercenaria mercenaria (Linne)was
sampled for trace metals to determine how trace metal con-
centrations in the shellfish tissues compared with the level
of bacteriological pollution in the growing water and the
NSSP classification. Hard clams were sampled from 5 loca-
tions in each harbor and analyzed for copper, lead, zinc, and
cadmium. From the metal and conform concentrations and
their distributions in the two harbors, the following relation-
ships were observed: in both harbors, hard clams from the
station with the fewest coliform bacteria did not have the
lowest metal concentrations; in Setauket, the variability in
metal concentrations among the sampling locations was
much less than in Port Jefferson; and in Port Jefferson, over-
all metal concentrations were higher than in Setauket. The
concentration of metals in the shellfish does not appear to
be reliably related to the coliform level.

BLUE CRAB PREDATION ON INFAUNAL BIVALVES:
RELATION TO OPTIMAL FORAGING HYPOTHESES

VICTOR S. KENNEDY, C. KING AND
J. BLUNDON

Horn Point Environmental Laboratories
University of Maryland. Box 775
Cambridge, Maryland 21 613

Adult blue crabs {Callinectes sapidus Rathbun) were
allowed to forage on equal numbers of 3 size classes of
buried soft-shell clams (Mya arenaria Linnd); percentage
of clams ingested increased with increasing clam size. This
was also true in the case of juvenile blue crabs foraging on
equal numbers of 5 size classes of buried specimens of
Macoma balthica (Linne). When the largest size class of M.
balthica was not available and equal numbers of the four
remaining size classes could be preyed upon by juvenile
crabs, the percentage of clams ingested increased with
increasing clam size. This seems to indicate a pattern of
optimal foraging by the crabs. Equal biomass of (a) two size
classes of buried speimens of M. arenaria or (b) three size
classes of buried specimens of M. balthica was then made
availabe to adult or juvenile blue crabs, respectively. At the
end of these experiments there was no statistically significant
difference among size classes in percentage of clams ingested.
This suggests that buried clams are preyed upon opportunis-

tically by blue crabs. The results of the experiments using
equal numbers of clams per class may have been influenced
by the possibility that larger clams have a greater chance than
smaller clams of being encountered by a sediment-probing
crab because of their larger size.

DEPARTMENT OF NATURAL RESOURCES AND

UNIVERSITY OF MARYLAND FORM NEW

COOPERATIVE SHELLFISH RESEARCH

UNIT AT CRISFIELD

GEORGE E. KRANTZ

University of Maryland Center for
Environmental and Estuarine Studies
Box 775, Cambridge, Maryland 21613

The University of Maryland's Marine Products Laboratory
located at Crisfield has become the site of a joint University/
Department of Natural Resources (DNR) program in shell-
fish management effective 1 January 1982. The new joint
research and management program will offer many advan-
tages to the state's seafood industry by combining research
and management functions in one unit as well as providing
for the transfer of new hatchery technology through demon-
strations of shellfish culture methods to watermen, seafood
processors, and other interested groups.

THREE INNOVATIVE TECHNIQUES THAT MADE MARYLAND
OYSTER HATCHERIES COST-EFFECTIVE

GEORGE E. KRANTZ, G. J. BAPTIST
AND D. W. MERITT

University of Maryland Center for
Environmental and Estuarine Studies
Cambridge, Maryland 21613

The combined use of 3 innovative techniques reduced
the size of the physical plant of a Maryland oyster hatchery
by 65% and reduced the labor by 55%. Tahitian Isochrysis,
an unidentified algal strain that has an optimal growth tem-
perature between 24 and 30°C, eliminated the need for a
temperature controlled algae culture room in the hatchery.
Algae cultures were grown at ambient room temperature
and stored in a '"concentrated paste" after dewatering in a
mechanical centrifuge. This technique permitted year round
operation of a small algae culture laboratory rather than an
intensive period of activity during the time of oyster larval
culture (June through August). Oyster spat were collected
directly from larval culture cones on a concrete-coated, wire
device which also served as a growing substrate until the spat
reached 2.5 to 3.5 cm. This growing device was transferred
directly from the larval cone into the natural environment
thereby eliminating the need for continuous flow of water
in the hatchery and the labor involved with cleaning vast
expanses of spat culture trays. Field trials of spat grown by

Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

these techniques will yield marketable oysters in the fall of
1983.

EFFECT OF PROCESSING ON STEROL AND FATTY ACID
COMPOSITION OF CRABMEAT

JUDITH KRZYNOWEK

National Marine Fisheries Service
Northeast Fisheries Center
Gloucester Laboratory
Gloucester, Massachusetts 01 930

The use of water or brine or mechanical stress for crab-
meat extraction and the freezing or further heating of
crabmeat for canning purposes are processing techniques
employed by the crabmeat industry. The impact of physical
and chemical processing is discussed relative to the effect
on the lipid portion of the meat (primarily on the sterol
and fatty acid composition). Specific processing techniques
to be discussed include: freezing, multiple freeze/thaw
cycles, canning (both sterilized and pasteurized and the
inclusion of bacteria in the product after canning), and three
methods for meat extraction.

ESTIMATION OF STANDING CROP OF MERCENARIA

MERCENARIA (LINNE) IN THE JAMES RIVER,

VIRGINIA, USING COMMERCIAL RECORDS

ANDRE C. KVATERNIK AND
WILLIAM D. DUPAUL

Sea Grant Marine Advisory Services
Virginia Institute of Marine Science
College of William and Mary
Gloucester Point, Virginia 23062

Commercial catch and effort records for boats using
patent tongs to harvest hard clams from the James River
were obtained for the years 1978-1981. Using Dickie's
(1955) version of the Leslie method, catch-per-unit-effort
of the sample fleet was regressed against accumulated catch
to give estimates of the initial abundance. Estimates for
1978, 1979, 1980, and 1981 were 10,101 m 3 (280,605 bu),
14,625 m 3 (406,250 bu), 20,065 m 3 (557,250 bu), 1 2,397 m 3
(344.364 bu), and 14,297 m 3 (397,142 bu), respectively. The
mean for the period 1978-1981, 14,297 m 3 (397,142 bu),
was 30% below that estimated by Haven et al. (1981). Com-
mercial catch records can be used in this application but
limitations in the data must be understood. Abundance esti-
mates from this method should be supplemented with addi-
tional designed sampling strategies to give better accuracy.

EFFECTS OF LIGHT AND GRAVITY UPON THE MOTILE

BEHAVIOR OF TROCHOPHORE LARVAE OF

MERCENARIA MERCENARIA (LINNE)

MARK D. LESLIE AND
ROBERT S. WILSON

Department of Biology

Southeastern Massachusetts University

North Dartmouth, Massachusetts 02747

Adults of Mercenaria mercenaria were spawned in the
laboratory and the fertilized eggs were reared to the trocho-
phore stage. Responses of the larvae to light and gravity were
observed. Distributions were determined under 5 experi-
mental conditions: horizontal chamber in darkness, horizon-
tal chamber with two different light intensities (2.5 and 15
W/M 2 ) shining from one end, vertical chamber in darkness,
vertical chamber with light incident from above (2.5 W/M 2 )
and a vertical chamber with light incident from below (2.5
W/M 2 ). The results revealed a random distribution of the
larvae in horizontal dark and horizontal light experiments, a
substantial surface aggregation in the vertical dark chamber,
and a decrease in surface accumulation with the light source
shining from above and below the vertical chamber. Indivi-
dual swimming paths of the larvae were analyzed using a
computer-video system (viz., the Bug-system). The larvae
were viewed in both the presence and absence of light in a
vertical plane. Illumination from below caused a significant
drop in vertical velocity and swimming speed and a small
decline in the rate of change of direction. Phototaxis was
not observed. Photostimulation caused the trochophores to
exhibit a negative orthokinesis with a weakening in their
negative geotactic behavior.

GROWTH OF JUVENILES OF ARCTICA ISLANDICA (LINNE)
IN EXPERIMENTAL CONTAINERS

R. A. LUTZ 1 , J. G. GOODSELL 1 ,
M.CASTAGNA 2 AND A.P.STICKNEY 3

Dept. of Oyster Culture, New Jersey
Agricultural Experiment Station, Cook
College. Rutgers University
New Brunswick, New Jersey 08903

Virginia Institute of Marine Science
Wachapreague, Virginia 23480

Dept. of Marine Resources
West Boothbay Harbor, Maine 04575

Laboratory -reared ocean quahogs {Arctica islandica)
(n = 119) ranging in shell length (maximum antero-posterior
dimension) from 1.8 to 4.3 mm (x = 2.5 ± 0.4 mm, SD)
were placed during June in experimental mesh containers
suspended from fixed and floating structures in marine
waters off Boothbay Harbor, Maine. Shell length measure-
ments were recorded at monthly intervals until the follow-
ing March. Water temperatures at the locations of the con-
tainers ranged from a high of 15.5°C during August to a
low of 1 .0°C during February. Mean growth rages recorded
during the warmer months from June through September

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

ranged from 2.0 to 2.4 mm/month. Reduced, yet measur-
able, amounts of shell (x = 0.3 - 0.5 mm/month) were
deposited during even the coldest winter months (January
and February). Mortality during the study period was < 1%.
By early March, the shell lengths of specimens (n = 117)
ranged from 3.9 to 21.3 mm (x = 14.0 ± 2.8 mm, SD).
Recorded growth rates were considerably faster than those
heretofore reported for Arctica islandica and suggest that
juveniles of this species have a potential for relatively rapid
growth in certain environments.

SIZE AND VOLUME RELATIONSHIPS IN JUVENILES OF

MERCENARIA MERCENARIA (LINNE):

A REVISION OF BELDINGS TABLES

JOHN J. MANZI ' , F. S. STEVENS 1 ,
Y. M. BOBO 1 , V. G. BURRELL, JR. 1
AND NANCY H. HADLEY 2

Marine Resources Research Institute
Charleston, South Carolina 29412

College of Charleston,
Charleston, South Carolina 29402

Size and volume relationships in juveniles of the hard
clam Mercenaria mercenaria were determined in commer-
cial nursery populations over a 1-y period. Morphometric
determinations included size (longest anterior-posterior
dimension), displacement volume, and packed volume (wet).
These data were used to establish empirical relationships
between seed size and volume (displacement and wet
packed) which are reported here as a revision of Belding's
Tables. The empirical relationships, thus established, were
iteratively employed in the construction of a model to pre-
dict seed clam volume. The model assumed that the volume
of a hard clam is proportional to the cube of a linear dimen-
sion. The iterations allowed model refinements which pro-
duced positive correlations between predicted and observed
data. We summarize collected data on size/volume relation-
ships in seed clams and present a model, based on truncated
spheres, which attempts to relate size and volume character-
istics in seed clams within the size range of 1 .0 to 1 5 .0 mm.

A DESCRIPTIVE MODEL FOR THE CONSERVATION OF

BLUE CRAB LARVAE IN THE VICINITY OF

CHESAPEAKE BAY

J. R. McCONAUGHA, D. R. JOHNSON
AND A. J. PROVENZANO

Department of Oceanography
Old Dominion University
Norfolk, Virginia 23508

An extensive series of plankton samples taken from the
waters around Chesapeake Bay indicates that all larval stages
of the blue crab Callineetes sapidus Rathbun are concentra-
ted in the upper layers of the water column with maximum

numbers in the upper 1 m. This distribution insures that
stage I larvae hatched near the bay mouth are entrained in
the outwardly flowing surface water. The general longshore
current in the Mid-Atlantic Bight is southward which would
tend to transport larvae towards Cape Hatteras. This would
result in their being lost to the system. Recent evidence sug-
gests that during the summer months, when peak spawning
occurs, there is a wind generated counter-current on the
inner shelf. The width and speed of this corridor is related
to wind direction and velocity. Larvae entrained in this
counter-current are returned to the vicinity of Chesapeake
Bay and contribute to recruitment. The horizontal distri-
bution of blue crab larvae from field samples is consistent
with this hypothesis.

A TEST OF A DART TAG FOR JUVENILE BLUE CRABS,
CALLINECTES SAPIDUS RATHBUN

R. E. MILLER

University of Maryland

Horn Point Environmental Laboratories

Cambridge, Maryland 21613

A small dart tag was applied to the posterior junction
between the ventral and dorsal parts of the cephalothorax
of 80 juvenile blue crabs to test for success of molting and
tag retention during the molting process. Sixty-one percent
of tagged crabs which began ecdysis were successful in
molting and retained the tag; however, overall mortality
rate for tagged crabs was twice that of the untagged control
group.

METHODS FOR FIELD EXPERIMENTS
WITH BAITED TRAPS

ROBERT J. MILLER

Fisheries Research Branch, Canada
Department of Fisheries and Oceans
Halifax, Nova Scotia, Canada, B3J2S7

The number of uncontrolled variables and the number of
potentially testable variables in the field environment can
be distracting and intimidating to the field technicians. This
environmental complexity requires greater mental discipline
to conduct good experiments in the field than is required in
the tidier laboratory environment. Problems frequently
encountered in conducting experiments on design and fishing
strategy of baited traps are as follows. Testing of hypotheses
using fishermen's logbook data commonly gives biased
results and has poor resolution because fishing variables are
neither controlled nor random and data are often incorrect.
Because most fishermen lack appreciation for correct experi-
mental procedures, even dictating an experimental design
will not assure a properly executed experiment. Preliminary

Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

trapping should be carried out to locate an experimental area
with uniform catch rates, to determine the optimum sample
size, and to solve logistical problems in conducting the
experiment. Experimental treatments should be randomized
in space and time to avoid bias. An investigator rarely knows
enough about the uncontrolled variables in the field to jus-
tify a systematic allocation of treatments in space and time.
Variance is controlled by careful attention to details of bait
quantity and quality, by keeping traps in good repair, by
standardizing soak time, and by standardizing time of day
of setting traps.

A FIRST ESTIMATE OF INDIRECT FISHING

MORTALITY IN THE ICELAND SCALLOP

CHLAMYS ISLANDICA (MU LLER)

K. S. NAIDU

Research and Resource Services
Department of Fisheries and Oceans
P.O. Box 5667. St. John's.
Newfoundland, Canada A1C 5X1

Natural mortality in Iceland scallops (Chlamys islandica),
computed from the ratio of cluckers to live animals, as
might be expected, increased with age. Higher than average
rates were found for the fully recruited ages (> 8 y) on
heavily exploited grounds than in scallop beds subject to
light or initial exploitation. The difference in mortality rates
between near-virgin and fully exploited areas is ascribed to
indirect fishing mortality associated with repetitive towing
on productive grounds.

THE ANNUAL GLYCOGEN CYCLE IN THE SOFT-SHELL CLAM
MYA ARENARIA LINNE FROM MAINE

CARTER R. NEWELL

Program in Oceanography . University of
Maine at Orono, Ira C. Darling Center
Walpole, Maine 045 73

A field population of adults of Myaarenaria was sampled
at approximately semi-monthly intervals for one year to
determine glycogen levels in the meats. Highest levels
occurred in late spring and early summer. Post-spawning late
summer and fall levels were intermediate, and lowest levels
occurred in the winter. Glycogen levels in juveniles and adults
of M. arenaria were compared and the relationships between
glycogen levels and gametogenesis, food availability, and
temperature are discussed.

THE EFFECTS OF SEDIMENT TYPE ON GROWTH RATE

AND SHELL ALLOMETRY IN THE SOFT-SHELL CLAM

MYA ARENARIA LINNE

CARTER R. NEWELL

Program in Oceanography, University of
Maine at Orono, Ira C. Darling Center
Walpole. Maine 04573

Hatchery-reared juveniles of Mya arenaria were grown
for 1 1 weeks in replicated gravel, sand, mud, and pearl net
treatments under flow-through seawater conditions in Maine.
Analyses of variance showed significant differences between
sediment treatments for final shell length, dry meat weight,
chondrophore growth increment, and percent shell weight.
Growth of juveniles of M. arenaria was more rapid in fine
sediments than in coarse sediments or nets. The slopes of
shell length vs. shell height and shell length vs. shell depth
regressions also varied significantly between sediment treat-
ments. Slower growing clams from nets and gravel were more
globose than clams from sand or mud treatments. Clams
reared in sand were longer and narrower than those reared
in mud. Differences in growth rates and shell form were
attributed primarily to the physical properties of the sub-
stratum.

PREFERENTIAL INGESTION OF ORGANIC MATERIAL FROM

THE CONSUMED RATION BY THE OYSTER

CRASSOSTREA VIRGINICA (GMELIN)

ROGER I. E. NEWELL AND
STEPHEN JORDAN

Horn Point Environmental Laboratories
University of Maryland. P.O. Box 775
Cambridge. Maryland 21613

Considerable debate exists in the literature as to whether
suspension-feeding bivalve molluscs can preferentially ingest
the organic component of the seston. Most of those discus-
sions were based on circumstantial evidence rather than reli-
able, quantitative measurements of the chemical composition
of the oyster's food or biodeposits. This paper gives details
of steady state measurements of the carbon, nitrogen, and
energy content of the seston being fed to the oyster Cras-
sostrea virginica and of the faeces and pseudofaeces being
voided. The results indicate that, over the tested range of
food concentrations (from 4—20 mg/1), the amount of
energy (expressed as Joules/mg of dry weight of material)
voided in the pseudofaeces by C. virginica can be reduced
by 60% compared to the concentration in the food. Similar
results were obtained from the carbon and nitrogen analysis.
These data strongly indicate that C. virginica has the capa-
bility of selecting certain particles from the total seston
filtered from suspension, with the result that more food
particles are rejected in the pseudofaeces.

FACTORS LIMITING ABUNDANCE OF
CALLINECTES SPP.

ELLIOTT A. NORSE 1 AND
VIRGINIA FOX-NORSE 2

Center for Environmental Education
624 9th Street NW,
Washington. D.C. 20001

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982

United States Environmental
Protection Agency, Office of Federal
Activities, A- 104
Washington, DC 20460

The abundance of organisms varies in space and time be-
cause the factors that limit abundance vary spatially and
temporally. Understanding limiting factors and the ways
organisms respond to them can lead to improved blue crab
catches. Blue crab populations can be limited directly by
(1) insufficient recruitment from the plankton; (2) inade-
quate water quality, due either to natural or man-made causes;
(3) insufficient resources, including food and cover;(4) inter-
ference competition, especially from other crabs; and (5)
removal by parasites, natural predators, and crabbers. Each of
these classes of limiting factors can be tested experimentally.
The results of these studies can suggest more effective ways
to improve catches by managing not only the populations
of blue crabs, but also the ecosystems to which they belong.

TOTAL WIDTH -WEIGHT RELATIONSHIPS OF THE BLUE

CRAB CALLINECTES SAPIDUS RATHBUN FROM THE

ASHLEY RIVER, SOUTH CAROLINA

EUGENE J. OLMI, III AND
JAMES M. BISHOP

Marine Resources Research Institute
South Carolina Wildlife and Marine
Resources Department
Charleston, South Carolina 29412

Equations expressing total width-weight relationships of
blue crabs (Callinectes sapidus) were calculated in relation
to sex, sex by maturity, sex by molt sign, and sex by cara-
pace form. All calculations were restricted to intermolt
(Stage C) crabs except when molt sign was considered, and
comparisons were restricted to crabs of similar size. Sex,
maturity, molt sign, and carapace form significantly affected
width-weight relationships. Overall, males were heavier than
females of equal width. Mature males exhibited a greater
mean weight than immature males, but mature females
weighed less than immature females of similar size. Crabs
with short lateral spines were heavier than those of the
same sex with long spines. Intermolt and premolt (Stage D)
males and females were heavier than recently molted (Stages
A and B) males and females, respectively. Premolt females
were heavier than intermolt females; a similar difference
was not observed for males. Ashley River crabs were generally
heavier than crabs from Florida, Texas, and Virginia. These
differences may not be real, however, because many variables
affect width-weight relationships of blue crabs and only sex
differences were reported. Geographical variation is known
to exist in crab populations, but only well defined compari-
sons between populations should be considered.

SIGNIFICANCE OF THE NEUSTON LAYER IN THE

DISPERSAL OF LARVAE OF THE BLUE CRAB

CALLINECTES SAPIDUS RATHBUN

A.J.PROVENZANO,J.M.
McCONAUGHA, AND D.F. JOHNSON

Department of Oceanography
Old Dominion University
Norfolk, Virginia 23508

The distribution of larval blue crabs in the water column
affects their transport out of Chesapeake Bay and during
the larval period. The patterns of vertical distribution are
not similar to those of other crab species in the region. First
stage larvae are found predominantly in the neuston layer
during the hatching season in the mouth of Chesapeake Bay
and are carried seaward by the ebb tides. Later develop-
mental stages, including the megalopae, are also found pre-
dominantly in the neuston or upper 1 m, with very few being
caught in intermediate layers or near bottom. Up to 99% of
stage I larvae in the bay mouth and more than 70% of all
Callinectes larvae of all stages even offshore were found
above lm. No evidence of vertical migration of any stage
was obtained. The effect of this distribution is to make
larval blue crabs very susceptable to surface effects and wind
driven currents during larval development and immediately
after metamorphosis to the megalops. Studies which do not
include the neuston layer may overlook a major fraction of
the total population of blue crab larvae. Most previous
studies of larval blue crab occurrence and distribution did
not include sampling of the neuston and consequently some
conclusions based on those studies were erroneous.

GROWTH ENHANCEMENT OF MY A ARENARIA LINNE

AND MERCENARIA MERCENARIA (LINNE)

BY MARINE MACROALGAE

HAUKE K. RASK

Ira C. Darling Center, University of Maine
Walpole, Maine 04573

Juveniles of My a and Mercenaria were Alizarin-stained
and cultured for 12 weeks in flow-through tanks containing
one of three different species of macroalgae. Clams grown
with Ascophyllum nodosum Linnaeus and Laminaria longi-
cruris De la Pylaic were significantly larger with respect to
shell dimensions than controls and those grown with Ulva
lactuca Linnaeus. Maximum enhancement was observed
with Ascophyllum in all czses;Mya grown with Ascophyllum
grew 4.54 times more than controls, while Laminaria treated
My a showed 2.14 times more growth. A similar but less
pronounced trend was seen for Mercenaria. Treatments
with Ascophyllum and Laminaria were 12.6% and 9.6%
larger than controls, respectively. Growth with Ulva was less

100 Abstracts, 1982 Annual Meeting, June 14-17. 1982

National Shellfisheries Association, Baltimore, Maryland

than control treatments but differences were not significant.
The mechanisms of growth enhancement from different
macroalgae and their importance in aquaculture are discussed.

ECONOMIC CONSIDERATIONS IN MANAGEMENT OF THE
COMMERCIAL BLUE CRAB FISHERY

RAYMOND J. RHODES

Division of Marine Resources
South Carolina Wildlife and Marine
Resources Department
Charleston, South Carolina 29412

From an economic prospective, the major consideration
of common wealth fishery management is to maximize net
benefits derived from the resource. In the case of commer-
cial fisheries, net benefits accruing to society should include
harvest revenues minus private costs (e.g., public adminis-
tration and enforcement). In order to accomplish manage-
ment objectives, private costs and public transactions costs
need to be minimized. A simple review of various blue crab
regulations germane to these economic concepts was
performed.

CHEMICAL ECOLOGY OF OYSTER DRILLS

M. CARRIKER 1 , L. WILLIAMS\ AND
L. WOOD 3

University of Delaware, College of
Marine Studies, 700 Pilot town Road
Lewes, Delaware 1 9958

Department of Botany. University of
Washington, Seattle, Washington 98195
3 101 Whitcomb Circle
Lafayette, Louisiana 70503

Oyster Drills are predatory snails that eat a wide spectrum
of shelled prey such as oysters, mussels, and barnacles. Drills
have a well documented ability to locate intact prey from a
distance by following chemical trails. We have looked in
detail at the molecular basis of prey location by drills. Newly
hatched drills can locate only barnacles from a distance. This
ability is apparently genetic as maternal diet and prey odor
environment do not enable the young to locate other prey
such as oysters or mussels. Once a newly hatched drill has
fed for some time on oysters, however, it develops the
ability to locate oysters. The molecules used by drills to
locate either barnacles or oysters are similar peptides.
Animals that can locate only barnacles, however, cannot
use even high concentrations of oyster attractant to locate
oysters. Drills cannot locate mussels from a distance even if
they have fed upon mussels. In fact, mussels produce a
molecule that suppresses the ability of drills to locate prey
from a distance. This molecule is much different than the
attractant molecules. It has a molecular weight less than

500 Daltons and does not appear to be a peptide. As a result
of the differences between attractants and suppressants and
the responses of inexperienced versus experienced drills we
can measure levels of attractants and suppressants in natural
waters. We hope that an understanding of the molecules
and mechanisms involved in prey location can provide a
means of drill control in the near future.

DOCUMENTATION OF ANNUAL GROWTH LINES IN THE
OCEAN QUAHOG.4/?77C4 ISLANDICA LINNE

J. VV. ROPES 1 , D. S. JONES 2 , S.A.
MURAWSKl',F.M.SERCHUK 1 AND
A. JEARLD, JR. 1

U.S. Dept. Commerce. NMFS
Woods Hole, Maine 02543

Dept. Geoi, Univ. Florida
Gainesville, Florida 32611

About 42,000 ocean quahogs (Artica islandica) were
marked for release at a deep (53-m) oceanic site off Long
Island, NY, in 1978. Shells of live specimens recovered 1
and 2 years later have been radially sectioned, polished, and
etched for preparation of acetate peels and examination by
optical microscopy or microprojection; selected specimens
were similarly prepared for examination by scanning elec-
tron microscopy. Specific growth-line and growth-incre-
ment microstructures are described and photo-illustrated.
An annual periodicity of microstructure is documented.
The observations form a basis for resource assessment ageing
studies of the commercially important species.

THE CHESAPEAKE BAY BLUE CRAB FISHERY:
HISTORICAL TRENDS AND EMERGING ISSUES

LEONARD A. SHABMAN AND
TAMARA VANCE

Department of Agricultural Economics
Virginia Polytechnic Institute
Blacksburg, Virginia 24061

Twenty-year trends in the Chesapeake Bay (Virginia and
Maryland) blue crab fishery were measured with National
Marine Fisheries Service data. Despite a recent downward
trend in landings, Virginia continues to have the largest
annual harvest of blue crabs in the U.S. While the total
number of crabbers in Virginia has been stable, there have
been decreasing numbers of users of trotlines and dredges
and increases in users of pots. The mean harvest per crabber
has fluctuated with a perceptible downward trend; but con-
sistently rising ex-vessel prices have maintained rising gross
income in the fishery. Maryland landings, like Virginia's, are
a significant portion of U.S. harvest and have shown a slight
downward trend. The number of Maryland crabbers has
more than tripled over the observed period— predominantly
from additions to the recorded number of part-time laborers.

National Shellfisheries Association, Baltimore, Maryland

Abstracts, 1982 Annual Meeting, June 14-17, 1982 101

There has not been a decline in the use of trotlines in Mary-
land, as in Virginia, because of restrictions on the use of pots
in certain Maryland waters. In Maryland, the mean harvest
per crabber has fallen over the period. Consistently rising
ex-vessel prices have resulted in an upward trend in mean
labor income for Maryland pot crabbers, but there has been
a drop in mean labor income for Maryland trotline crabbers.
Based upon this review, three factors affecting the future
growth of the industry are discussed: (1) state laws to protect
brood stocks differ and confuse stock management efforts;
(2) current public management programs(primarily licensing)
may not be promoting maximum economic yield from the
fishery ;and (3) economic uncertainties restrain development
of processing facilities and, in turn, discourage harvest.

MANAGEMENT OF THE BLUE CRAB FISHERIES IN
NORTH CAROLINA: A CASE HISTORY

TERRY M. SHOLAR

North Carolina Division of Marine
Fisheries, Washington,
North Carolina 27889

Blue crabs support one of North Carolina's most impor-
tant fisheries. The recent expansion of the crab fisheries has
resulted in numerous management problems concerning
resource allocations and gear conflicts. Regulatory authority
for management in North Carolina has been delegated by
the General Assembly to a 15-member commission which
enacts regulations based on staff recommendations and
input from the industry and general public. A key manage-
ment tool is the proclamation authority which has been
delegated by the Commission to the Secretary of the De-
partment of Natural Resources and Community Develop-
ment to respond rapidly to management needs. Proclama-
tions can be issued to invoke a management action with a
minimum of 48 h of public notice. This is generally done to
open or close areas to a particular fishing method or to set
seasons. This ability allows effective response to rapidly
changing situations within the fisheries and the stocks. An
example of North Carolina's management system involving
the blue crab fisheries concerns resource allocation in certain
tributaries of Pamlico Sound. Potting and trawling are in-
compatible gears competing for space and resource. Each is
controlled by proclamation. The decision to allow a certain
fishery to occur is based on biological, economic, and social
implications, with multiple-use resource management and
protection being major factors in the decision. Tagging
studies are being used to evaluate management strategies
and their effect on maximizing crab harvest, and to deter-
mine short-term migratory habits. Numerous other manage-
ment issues affecting blue crabs and their fisheries such as
minimum size limit, mandatory cull rings in pots, spawning

sanctuaries, and nursery area protection are addressed.

THE TEXAS OYSTER STUDY. I. RELATIONSHIPS

BETWEEN AVAILABLE FOOD, OYSTER

COMPOSITION, CONDITION, AND

REPRODUCTIVE STATE

THOMAS M. SONIAT 1 AND
SAMMY M. RAY 2

Department of Biological Sciences
University of New Orleans, Lakefront
New Orleans, Louisiana 70148

Department of Marine Biology
Texas A &M University at Galveston
P.O. Box 1675, Galveston, Texas 77553

We examined the relationships between what is available
for the oyster to eat, the oyster's proximate composition.
its condition, and its reproductive state. Changes in the
proximate composition of oysters were associated with
changes in the annual cycle of fattening, storage, and repro-
duction. The fattening phase was characterized by high dry-
weight condition indices and elevated carbohydrate (glyco-
gen) concentrations. A "storage cycle," the transition from
stored glycogen to the lipid reserves in developing eggs, was
evident in Crassostrea virginica (Gmelin). The gonadal index
and percent lipid composition of the oyster were positively
correlated. Spawned oysters had low lipid and carbohydrate
concentrations, low condition and gonadal indices as well as
high concentrations of water and protein. Available food
for the oyster was measured as a food index. The food
index was defined as the percentage food (food = lipid +
carbohydrate + protein) in the total seston. The food index
was higher in the spring and summer and was correlated
with the gonadal index of oysters. Apparently, the amount
of food was greatest at the time of greatest energy demand;
that is, during gametogenesis.

THE TEXAS OYSTER STUDY. II. MODELS OF OYSTER
NUTRITION IN THE NATURAL ENFIRONMENT

THOMAS M. SONIAT 1 . SAMMY M.
RAY 2 AND REZENAT M. DARNELL 3

Department of Biological Sciences
University of New Orleans. Lakefront
New Orleans, Louisiana 70148

Department of Marine Biology
Texas A &M University at Galveston
P.O. Box 1675, Galveston, Texas 77553

Department of Oceanography
Texas A&M University
College Station, Texas 77843

Two FORTRAN models were developed to integrate in-
formation about measured food levels (i.e., the food index)
with the presumed needs of the oyster. One model assumed
no selective ingestion on the part of the oyster. Another
model assumed that the oyster could selectively ingest

102 Abstracts. 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

organic material. Although the results of the models are in
fair agreement with published literature, this agreement
could simply be fortuitous. The correspondence between
the models we developed and other works, however, suggests
the possibility that the food index is a useful measure of
available food, that the simplifications made in the models
are reasonable ones, and that enough particulate food was
present to sustain oysters in the area studied.

A CYTOGENETIC METHOD AS A TOOL FOR ASSESSING
THE CONDITION OF SHELLFISH LARVAE

S. STILES AND J. CHOROMANSKI

National Marine Fisheries Service
Northeast Fisheries Center
Milford Laboratory
Milford. Connecticut 06460-6499

As a means of assessing their condition at the cellular
level, cultured oyster larvae were examined cytologically by
employing a relatively simple squash technique. Chromo-
some groups, and normal and abnormal cells and nuclei
were evident. Bacteria also were discernible with this method.
These observations were an indication of general health of
the larvae in culture and provided some information regard-
ing subsequent development and survival. In addition to
being able to observe pathological states of the cells and
bacterial infections, one could use the procedure to deter-
mine the numbers of cells in mitosis as an indicator of
growth rate. Larvae, potentially, could be pre-treated with
colchicine to arrest cells in mitosis for counting the chromo-
somes to obtain karyotypes as an aid in plankton identifica-
tion. Cytological analyses of the larvae could have many uses
in toxicological studies, including bioassays, as well as in
hatchery rearing and breeding.

ISOLATION AND PARTIAL CHARACTERIZATION

OF A MALATE DEHYDROGENASE FROM

CRASSOSTREA VIRGINICA (GMELIN)

MARY L. SWIFT AND
S. LAKSHMANAN

Chemistry Department, University of
Maryland, College Park, Maryland 20742

Final details of glucose metabolism in marine bivalve
molluscs are yet to be elucidated. Malate dehydrogenase
(E.C.I .1 .1.37) has been implicated in the catabolic path-
way leading to the formation of succinate, a major end pro-
duct of anaerobic metabolism in bivalve molluscs. Further
clarification of this metabolic scheme may be gained by an
examination of the properties of malate dehydrogenase
(MDH). Homogenates of tissues of Crassostrea virginica
contain at least 3 MDH isoenzymes. One of these was iso-
lated from acetone powders of mantle and gill tissue by

ammonium sulfate fractionation, gel permeation, and ion-
exchange chromatography. Some properties of this prepara-
tion were determined. The Michaelis-Menten constants were :
K m(OAA) = 1.18xlO- 4 M; K m(NADH ) = 4.86xl(r 5 M;
Km(mal) = 1.35xlO*M; K m(NAD) = 1.30xlO" 4 M. The
following were not substrates: NADP + , a-ketobutyrate, a-
ketovalerate, a -ketoglutarate.D-malate, pyruvate, succinate,
oxomalonate. Tartronate, D-, L-, and mesotartrate were not
substrates and were found to be competitive inhibitors of
malate oxidation. The pH optima were: 7.6 for NADH oxi-
dation and 9.5 for NAD + reduction. MDH was inhibited by
p-chloromercuribenzoate and N-ethylmaleimide. Listed in
decreasing order of effectiveness, Cd ++ , Zn ++ , Cu ++ , Co ++
and Ni ++ inhibited NADH oxidation by MDH.

COMPARISON OF THE GROWTH OF CRASSOSTREA

VIRGINICA (GMELIN) AT FIVE ALGAL RATION

LEVELS WITH SPECIFIC REFERENCE TO

PREDICTIVE FEEDING EQUATIONS

EDWARD R. URBAN AND
G. D. PRUDER

College of Marine Studies
University of Delaware
Lewes, Delaware 19958

Mixtures of the algae Thalassiosira pseudonana Hasle et
Heimdal (clone 3H) and Isochrysis aff. galbana Parke (T-
ISO) were fed at each of 5 levels to juveniles of Crassostrea
virginica. The oysters were grown for 3 wk at 25 °C and a
salinity of 30 ppt. The relationship between algal ration level
and oyster growth is presented. The results are discussed
with specific reference to several feeding equations
either published or in use or both. Recommended algal
ration levels are compared for their relative effectiveness.
We show that neither cell number, nor volume, nor weight
constitute an acceptable parameter for comparing algal
species or bivalve species. We recommend that feeding
studies be carried out for any new combinations of algae
until the nutritive value of the algal species can be correlated
with physical characteristics and environmental conditions.
The prudent use of predictive algal ration equations as
management tools is discussed.

A BLUE CRAB MANAGEMENT PLAN:
OBJECTIVES AND RESPONSIBILITIES

W1LLARD A. VAN ENGEL

Virginia Institute of Marine Science
The College of William and Mary
Gloucester Point, Virginia 23062

The blue crab, Callinectes sapidus Rathbun. of the
Atlantic and Gulf coasts supports one of the major marine
fisheries of the United States. Regulatory authority

National Shellfisheries Association, Baltimore, Maryland

Abstracts. 1982 Annual Meeting, June 14-17, 1982 103

concerning licensing, size and sex limits, quotas, seasons,
gear restrictions, and other controls over harvesting within
its territorial waters rests with each state, retained by the
respective state legislatures, but may be delegated to a com-
mission. Regulations should be based on the best biological,
economic, sociological, and environmental knowledge and
provide for optimum yield from the resource. The blue crab
industry's problems are not limited to regulation of the har-
vest. They also include the need for federal and state assis-
tance in processing, marketing, and research; conservation
of the blue crab habitat; and an adequate data base. A com-
prehensive blue crab management program should protect
the resource, encourage and assist fishing with a minimum
of regulations, and promote utilization of the product.

THE BEHAVIORAL BASIS OF LARVAL DISPERSAL

AND RECRUITMENT IN THE BLUE CRAB

CALLINECTES SAPIDUS RATHBUN

W. F. VAN HEUKELEM AND
S. D. SULKIN

Horn Point Environmental Laboratories
University of Maryland
Cambridge. Maryland 21613

Laboratory experiments have demonstrated that Stage I
blue crab zoeae exhibit a number of behavioral traits which
should result in distribution high in the water column. These
traits include: negative geotaxis which is unaffected by salin-
ity changes of 5 ppt; high barokiness at hydrostatic pressures
exceeding 1 atmosphere; increased swimming rate with
increased salinity; positive phototaxis at light intensities of
> 1CT 3 W/m 2 ; maintenance of swimming speed with
decreasing temperature ;and the ability to traverse haloclines
of 10 ppt as well as sharp thermoclines. Because it is known
that female blue crabs migrate to the mouth of Chesapeake
Bay to spawn, these behavioral traits should result in massive
export of virtually all Stage I zoeae in surface waters. Field
evidence by other workers supports this contention. Mega-
lopae possess behavioral traits that differ from late zoeal
stages, chiefly, a highly sensitive pressure response, faster
swimming speeds, negative geotaxis, and possibly locomotor
rhythms that may enhance their transport back into estu-
aries. Since larval development occurs on the continental
shelf, recruitment success of megalopae back into estuaries
is likely to be highly dependent on offshore climatological
events that determine coastal circulation patterns during
the summer and fall.

REPRODUCTIVE PERIODICITY OF BUSYCON CARICA
(GMELIN) IN WATERS OFF SOUTH CAROLINA

DEBRA A. WEINHEIMER

Department of Biology
College of Charleston
Charleston, South Carolina 29424

A total of 1237 knobbed whelks (Busy con carica) were
collected over a 13-month period near Charleston Harbor,
SC. Gonad maturation stages were determined by gonad
color and histological sectioning. Monthly fluctuations in
gonad weight, penis or nidamental gland weight, gonadal
index, and reproductive index were also examined, of the
six reproductive characteristics used in this study, gonadal
index values were considered to be the best indicators of
periodicity. The highest gonadal index values for males
occurred in September, October, November 1979, and in
March 1980. The highest values for females occurred from
September 1979 through May 1980. Sex ratios fluctuated
monthly. The number of females was significantly higher
than the number of males from July 1979 through January
1980. This situation was reversed in April 1980 when the
number of males was significantly higher than the number of
females. Sex ratios also fluctuated when examined using
shell-length classes. The smallest individuals in the monthly
samples were females (60-64 mm). All individuals with shell-
length values > 159 mm were female. Sex ratio relationships
to reproductive periodicity are discussed.

DISTRIBUTION, SIZE, AND SEX COMPOSITION

OF THREE SPECIES OF CALLINECTES

IN THE COASTAL HABITAT OF THE

SOUTH ATLANTIC BIGHT

ELIZABETH L. WENNER AND
CHARLES A. WENNER

Marine Resources Research Institute
Charleston, South Carolina 29412

Collections by shrimp trawl during summer of 1980 at
depth of 4.5 — 18 m between Cape Fear, NC, and Cape
Canaveral, FL, showed that biomass of Callinectes sapidus
Rathbun was greater than that of the other 72 decapod
species collected. Callinectes similis Williams ranked fourth
in abundance among the other decapod species collected,
but C sapidus and C. ornatus Ordway were not as numerous.
Catches of Callinectes spp. were greatest in the nearshore
depth zone of 4.5— 8.5 m. Density and biomass totaled for
all strata were greatest for C similis and C. ornatus off Flor-
ida, and for C. sapidus off South Carolina. Few mature or
ovigerous females of C. similis andC. ornatus were collected,
whereas most females of C. sapidus were either mature or
ovigerous. Significantly more females than males of C. sapi-
dus were collected. The ratio of M:F for other Callinectes
spp. varied with location. Sizes of crabs were not correlated
with depth or distance from shore.

104 Abstracts, 1982 Annual Meeting, June 14-17, 1982

National Shellfisheries Association, Baltimore, Maryland

NURSERY CULTURE OF THE BAY SCALLOP ARGOPECTEN

IRRADIANS IRRADIANS (LAMARCK) IN SUSPENDED

MESH ENCLOSURES

JAMES C. WIDMAN, EDWIN W.
RHODES AND P. A. BOYD

Milford Laboratory. Northeast Fisheries
Center, National Marine Fisheries Service
212 Rogers Avenue,
Milford, Connecticut 06460-6499

Suspended mesh enclosures with bottom areas of 0.1 m 2
were used to grow hatchery-reared bay scallops in Long
Island Sound in 1980 and 1981. The enclosures were con-
structed of 3- or 7-mm polyethylene mesh and were

anchored at a depth of 8 m and buoyed with styrofoam
floats. Scallops as small as 4.6 mm were successfully grown
to a size > 20 mm in the units. Acclimated scallops deployed
in the spring of 1981 at temperatures as low as 5°C survived
and subsequently grew normally as water temperatures
increased. Scallop densities between 250 and 1 5,000/m 2 were
tested in the enclosures, and although final shell height was
inversely related to density, substantial growth occurred at
all densities. Biovolumes of up to 3.9 C/enclosure were
obtained. Some comparisons between culture of small
scallops in mesh enclosures in Long Island Sound and in
raceways were made and both systems were useful for
nursery culture of this species.

Journal of Shellfish Research, Vol. 3, No. 1, 105-115, 1983.

ABSTRACTS OF TECHNICAL PAPERS

Presented at 1982 Annual Meeting

WEST COAST SECTION

NATIONAL SHELLFISHERIES ASSOCIATION

Olympia, Washington
September 10-11, 1982

Olympia, Washington, September 10-12, 1982 Abstracts, 1982 NSA West Coast Section Meeting 107

CONTENTS

Richard Albright

Population Structure and Production of the Amphipod Corophium salmonis

Stimpson in Grays Harbor, Washington 109

/. H. Beanie and J. Perdue

Progress in the Development of Resistance Against Summer Mortality through

Selective Breeding of Pacific Oysters 109

Clarke G. Beaudry

Survival and Growth of the Larvae of Haliotis kamtschatkana Jonas

at Different Temperatures 109

Richard Bumgarner

Recent Developments in the Spot Prawn Fishery in Hood Canal, Washington 110

Ken Cooper

Potential for Application of the Chemical DOPA to Commercial Bivalve

Setting Systems 110

Flinn Curren

Japanese Oyster Drill Studies Ill

Catherine Falmagne

Problems Associated with the Rearing and Setting of Larvae of the

California Mussel Mytilus californianus Conrad in a Hatchery 112

Jill E. Follett

A Histological Study of the Gastrointestinal Tract of the

Tanner Crab Chionoectes bairdi Rathbun (Decapoda, Reptantia) 112

Thomas C Kline

The Effect of Population Density on the Growth of the Butter Clam Saxidomus gigantus 112

Nancy Musgrove

The Feeding Behavior of the Terebellid Polychaete Thelepus crispus Johnson

in Response to Currents 113

Louisa Nishitani and Kenneth Chew

Vertical Migration of Gonyaulax catenella: Potential Implications for Management

of Paralytic Shellfish Poisoning (PSP) Problems 113

Scharleen Olsen

Abalone and Scallop Culture in Puget Sound 113

Timothy Sample

PSP: Its History, Processes and Impacts as Applicable to Puget Sound 114

A. Kimbrough Siewers

Commercial Mariculture of a Bay Scallop Argopecten circularis (Sowerby) in

the Ensenada of La Paz, Baja California Sur, Mexico 114

John J. Sullivan and Wayne T. Iwaoka

PSP Research: Recent Advances in Analytical and Biochemical Studies 114

Louis Wachsmuth

Disaster Ahead for the Yaquina Bay Oyster Industry? 115

Olympia. Washington. September 10-12, 1982

Abstracts, 1982 NSA West Coast Section Meeting 109

POPULATION STRUCTURE AND PRODUCTION OF THE

AMPHIPOD COROPHIUM SALMONIS STIMPSON IN

GRAYS HARBOR, WASHINGTON

RICHARD ALBRIGHT

Division of Aquaculture and
Invertebrate Fisheries. School of
Fisheries, University of Washington
Seattle, Washington 98195

The tube-dwelling amphipod Corphium salmonis is a
dominant benthic organism and important food resource in
the estuarine mudflats of Grays Harbor, WA. Intertidal core
samples were collected at two sites during the spring and
summer of 1980 to determine the population structure,
biomass, rate of growth, and production of C. salmonis.
The abundance of C. salmonis ranged from 200 to 50.000
individuals per m 2 . Peak abundances occurred during July
and August. Abundances at the 1 .8-m stations were higher
than at the 0.6-m stations. Females of C. salmonis attained
sexual maturity at alength of 4.0— 4.5 mm. Brooding of eggs
began in April and continued through the end of sampling
(30 September). Male-female ratios were lower for sexually
mature individuals of C salmonis than for immature individ-
uals, apparently as a result of predation on sexually mature
males which wander over the tideflats in search of females.
Male-female ratios decreased in the lower intertidal zone,
apparently as a result of increasing predation pressure.
Ratios also decreased over time at all stations, suggesting
that predation pressure may also increase through the spring
and summer. An inverse relationship between male-female
ratios for mature and immature amphipods suggests a pos-
sible genetic response to disparate sex ratios among mature
individuals. Data from both natural populations and from
cohorts which were artificially isolated inside in situ cages
were used to obtain size-specific growth rate curves and
production estimates for C. salmonis. Total Corophium pro-
duction for each station between 1 April and 30 September
varied from 3.6 to 10.7g/m 2 dry wt. Corophium production
was higher at the upper intertidal stations. Turnover rates
(the ratio of production to mean biomass) ranged rom 7.2
to 8.6. The production and turnover rates of Corophium
salmonis are high relative to other invertebrate species.
Thus, this amphipod is an important contributor to secondary
production in Pacific Northwest estuaries, providing an
important food resource for its predators, many of which
are commercially or recreationally valuable. This production
must be taken into consideration when making decisions
relating to activities such as dredging and filling which have
potentially adverse impacts on intertidal areas.

PROGRESS IN THE DEVELOPMENT OF RESISTANCE
AGAINST SUMMER MORTALITY THROUGH
SELECTIVE BREEDING OF PACIFIC OYSTERS

J.H. BEATTIE AND J. PERDUE

Division of Aquaculture and
Invertebrate Fisheries, School of
Fisheries, University of Washington
Seattle, Washington 98195

Since 1974 the University of Washington^ School of
Fisheries has been conducting research in the genetics of
the giant Pacific oyster Crassostrea gigas (Thurnberg). The
main emphasis of this work has been the development,
through selective breeding, of oyster stocks with high sur-
vival potential during summer mortality. Summer mortality
is a phenomenon that routinely accounts for losses of from
10 to 60% of harvestable 2-year-old oysters in bays of the
states of Washington and California, and Japan. The breed-
ing program began as a selection of individuals from wild
populations. The selection process was based upon survival
during elevated temperature (21 °C) challenges. The breed-
ing of these individuals (one male mated with one female)
produced families of oysters which could be tested and
compared on growing grounds experiencing annual mortal-
ities. On the basis of high survival during actual summer
mortality, families were selected as the brood lines for future
generations. Of 103 families tested since 1977, up to 78
have had higher survival than non-selected controls. The pri-
mary goal of the breeding program is to provide brood
stock to commercial hatcheries for production of oyster
seed resistant to summer mortality. However, for the past
three years, the families have also been monitored for
growth, gonadal development, and glycogen storage. Since
reduced gonadal development and high glycogen content
are desirable commercial characteristics, these parameters
have also been used in our overall breeding plan. Brood
stocks which appear to show promise have been made
available to commercial hatcheries since 1978. Data are
now being processed and evaluated from the experimental
families which will provide valuable information concerning
heritability of glycogen levels, and experiments are being
conducted on the effects of inbreeding. With every step, an
understanding of oyster genetics is clearer and the goal of
commercial production of superior oysters is closer.

SURVIVAL AND GROWTH OF THE LARVAE OF

HALIOTIS KAMTSCHATKANA JONAS

AT DIFFERENT TEMPERATURES

CLARKE G. BEAUDRY

Division of Aquaculture and
Invertebrate Fisheries. School of
Fisheries, University of Washington
Seattle, Washington 98195

110 Abstracts, 1982 NSA West Coast Section Meeting

Olympia, Washington. September 10-12, 1982

Larvae of the pinto (or threaded) abalone 7/a//or/s kamt-
schatkana were reared at four temperatures, 14, 16, 18.5,
and 21°C in 2-C glass beakers. Survival at the end of the
experimental period was best at 18.5° and worst at 21°. More
rapid settlement observed at higher temperatures may have
improved survival at those temperatures by shortening the
vulnerable planktonic stage during which most mortalities
occurred. Abalone at the highest temperature (21°) showed
signs of thermal stress and experienced total mortality.
During early embryonic development, from fertilized egg
through the trochophore, the lowest temperature (14°) pro-
duced the most normal larvae and highest survival. At higher
temperatures progressively more mortalities and abnormal-
ities occurred. Larvae reared at 18.5° were consistently of
greatest size at settlement; however, abalone reared at 16°
grew more rapidly and obtained the greatest length at the
end of a 2-month period.

RECENT DEVELOPMENTS IN THE SPOT PRAWN FISHERY
IN HOOD CANAL, WASHINGTON

RICHARD BUMGARNER

Washington Department of Fisheries
Point Whitney Shellfish Laboratory
Brinnon, Washington 98320

Hood Canal, a major arm of Puget Sound, is located in
northwestern Washington about 48 km (30 mi) west of
Seattle. This is the only area in Washington that has consis-
tently produced commercial quantities of the spot prawn
Pandalus platyceros Brandt. Harvest for both commercial
and personal use (recreation) has been restricted to shell-
fish pot gear since the early 1950's. Increased commercial
fishing pressure and poor recruitment between 1972 and
1974 resulted in a decline in spot prawn abundance and
serious conflict between commercial and recreational fisher-
men. This necessitated emergency season closures in 1974,
1975, and 1976. The year 1977 marked the beginning of a
new management approach for the Hood Canal spot prawn
stocks and associated fisheries. Season lengths and opening
dates were set according to the results of a preseason stock
assessment and anticipated fishing effort. To ensure an equi-
table share of the available surplus for recreational fishermen
the season was opened first to sport fishing and later to
commercial harvest. By 1979, all fishermen were restricted
to the use of shellfish pot gear having a mesh size of
> 2.2 cm (7/8 in). This was initiated to protect juvenile
prawns and to increase total yield. Changes in management
appear to be working well. Since 1977, stock abundance
has increased from a pre-season index of 1 .1 3 kg (2.5 lb) to
3.06 kg (6.75 lb) per pot in 1982. Harvest is also at an all
time high. Nearly 95 metric tons were taken in both 1981

and 1982. Improved fishing success has also, in part, led to
a tremendous increase in fishing pressure. The rate of
increase has averaged nearly 50% per year since 1977. Better
methods of effort-control are now needed to deal with the
rapid expansion of this fishery.

POTENTIAL FOR APPLICATION OF THE CHEMICAL DOPA
TO COMMERCIAL BIVALVE SETTING SYSTEMS

KEN COOPER

Department of Biology
Humboldt State University
Areata, California 95521

Simple chemical compounds have been shown to trigger
attachment and metamorphosis of the larvae of several
species of marine invertebrates. The simplest molecules in
which settlement inducing activity has been demonstrated
are L-3. 4-dihydroxyphenylalanine (DOPA), gamma-
aminobutyric acid (GABA), and choline. These molecules
occur in the marine environment as covalently bounded
compounds associated with adhesives, lubricants, exoskeletal
proteins, and pigments. A review of numerous studies clearly
implicated these chemical cues in successful habitat selec-
tion by invertebrate at the termination of the planktonic
stage of the life cycle. The similarity between these mole-
cules and neurotransmitters suggests that the chemoreceptors
are modified either ontogenetically or phylogenetically from
receptors specific to the neurotransmitters dopamine. GABA.
and acetylcholine. Selectivity in response by larvae to a
given chemical appears to depend on the neurotransmitter-
like portion of the compound, whereas specificity appears to
depend on the protein, carbohydrate, or lipid constituents.
Pediveligers of the blue mussel Mytilus edulis Linne and the
giant Pacific oyster Crassostrea gigas (Thurnberg) settle
in response to the amino acid DOPA. Implementing the use
of chemicals to commercial setting systems depends on
being able to either modify the chemoreceptors so that they
respond to an inexpensive and easily available chemical
and/or manipulating settlement behaviors. The initial
objectives of my study were to determine the response of
oyster larvae to DOPA, to examine the potential for applica-
tion to existing commercial setting systems, and to
determine the effect of several environmental factors on
the degree of response. Aliquots of hatchery -reared pedivel-
igers of C. gigas were tested for attachment in culture dishes
to both aged oyster shells and the smooth glass surface of
culture dishes. The pediveligers were reared at 34 ppt and
at 25 °C. Within individual tests, the settlement response by
the pediveligers was examined following exposure to DOPA
at 0.00001 M while varying the salinity (25 to 35 ppt) and

Olympia, Washington, September 10-12, 1982

Abstracts, 1982 NSA West Coast Section Meeting 1 1 1

temperature (20 to 30°C). Controls were run without the
addition of DOPA. The results presented are preliminary
findings and only indicate observed trends. In tests which
offered only a smooth glass surface for settlement, attach-
ment of the larvae to the glass occurred after 24 hr with but
not without the addition of DOPA to the seawater. In tests
to which DOPA was added the highest percentage of attach-
ment occurred at a salinity /temperature combination of 35
ppt/30°C. The pediveligers also attached to the glass surface
at the following salinity/temperature combinations listed in
order of decreasing percent response: 35 ppt/25°C, 35 ppt/
20°C. and 30 ppt/30°C. After 18 hr. a relatively high num-
ber of pediveligers attached to the glass surface in the runs
without DOPA at a salinity/temperature combination of 35
ppt/30°C. Also at 35 ppt/30°C in the runs with DOPA a
smaller, but significant, percentage of the pediveligers meta-
morphosed (indicated by new shell growth) without attach-
ing to the glass surface. This did not occur in any of the other
runs. The oyster pediveligers were next tested for attach-
ment to aged oyster shells in response to the addition
of DOPA. Preliminary results indicate that there was a
slightly greater set after 24 hr onto the shells in the tests
with DOPA. However, exposure of the larvae to DOPA also
promoted attachment to the glass surfaces of the culture
dishes. The consequence was that after 48 hr, the set onto
the shell was greater in the runs without DOPA, although
the total percentage of larvae which undergo metamorphosis
appeared to be the same. In the runs with DOPA a signifi-
cant percentage of the larvae either attached to the glass
surface or metamorphosed without attaching to any sub-
strate. These findings suggest that DOPA will not increase
the percentage of set onto oyster shells when the setting is
allowed to occur over several days. Rather, these findings
clearly suggest that the use of DOPA promotes extraneous
setting onto otherwise unfavorable substrates. However,
these findings do not discount the possibility that chemicals
can be used to obtain a more rapid set. The use of chemical
cues appeared more applicable to setting systems in which
no preferred setting substrate is used, such as in the setting
of clams and clutchless oysters.

JAPANESE OYSTER DRJLL STUDIES

FLINN CURREN

Division of Aquaculture and Invertebrate
Fisheries, School of Fisheries
University of Washington
Seattle, Washington 98195

The Japanese oyster drill Ocenebra inomata (Recluz) is
an economically important predator of oysters in areas along
the west coast, as well as in its native Japan. Since its acci-
dental introduction into Puget Sound with shipments of

Pacific oyster seed, attempts to control this snail have
included expensive hand picking and mercuric chloride.
These animals aggregate during certain times of the year, and

it is suspected that this behavior is cued by water-borne
pheromones (chemical substances which enable communica-
tion between animals). Pheromones are currently being used
in the control of several insects (e.g., gypsy moth) and might
have potential as a control technique for the Japanese oyster
drill. It was necessary, therefore, to develop an appropriate
bioassay to test different water extracts for pheromones.
Bioassays consist of subjects (in this case snails), stimuli
(water with suspected chemical agents), and responses
(which should be easy to identify . associated with the stimuli,
reproducible, and rapid). Bioassays should also minimize
the water used for stimulus and control to decrease efforts
involved in chemical extraction and concentration. Large
numbers of snails must be assayed to give statistical credi-
bility to sometimes subjective behavioral data. Several bio-
assays have been based on the snail's rheotactic response (in
a current of water, the snail moves upstream). The Pratt
choice chamber was rejected because large volumes of water
were needed with only one snail per run. Riffle flumes were
rejected because turbulent flows were encountered. Cephalic
antennal elongation (after pipetting a small amount of water
in front of the snail) was also rejected because of ( 1 ) the
highly subjective nature of the response (i.e., when are the
antennas elongated?), and (2) the large time requirement of
(10 min/subject) with the undivided attention of the
research. The inadequacies of these bioassays led to work
currently being done on a trough bioassay. A test chamber
1 X 1.5 m (39 X 50 in.) was constructed with stimuli and
controls (aged sea water) entering the flume through over-
flowing 1-2 beakers. Several hundred snails were placed 1 m
from the beakers and the numbers of snails climbing up the
beakers during a 6-hr period are noted. Current research
using this apparatus includes: (1 ) dye studies to determine
the water depth necessary for good mixing; (2) determina-
tion of the threshold flow rate to induce rheotaxis in oyster
drills; (3) testing of flow rates with a known stimulus
(oyster effluent); and (4) testing of stimuli from whole
ground snail extracts and field-filtered effluents from aggre-
gations. Stimuli found to be effective in these bioassays may
eventually be used to bait traps or disrupt snail behavior to
control Japanese oyster drills on oyster beds.

112 Abstracts, 1982 NSA West Coast Section Meeting

Olympia, Washington, September 10-12, 1982

PROBLEMS ASSOCIATED WITH THE REARING AND

SETTING OF LARVAE OF THE CALIFORNIA

MUSSEL MYTILUS CALIFORNIANUS

CONRAD IN A HATCHERY

CATHERINE FALMAGNE

Division of Aquaculture and Invertebrate
Fisheries, School of Fisheries
University of Washington
Seattle, Washington 98195

Mytilus californianus was successfully spawned and its
larvae were reared through metamorphosis in the University
of Washington hatchery at Manchester, WA. Although suc-
cess in spawning and rearing may vary with the hatchery
location and methods, data indicated the unreliability of
induced spawning at any given time. Some effects resulting
from different experimental combinations of temperature
and salinity have been observed. Survival of larvae to the
pediveliger stage at 18°C and 32 ppt was 31%. The larvae all
settled at the lower part of the suspended seed ropes because
they have a tendency to sink to the bottom of the tank
throughout metamorphosis. Further, higher numbers of the
larvae settled when the water was "conditioned" with adult
mussels.

A HISTOLOGICAL STUDY OF THE GASTROINTESTINAL
TRACT OF THE TANNER CRAB CHIONOECTES
BAIRDI RATHBUN (DECAPODA, REPTANTIA)

JILL E. FOLLETT

Alaska Dept. of Fish and Game
333 Raspberry Rd.
Anchorage, Alaska 99502

The tanner crab Chionoecetes bairdi is a commercially
important species in Alaska about which little is known of
its histology. In this study of the tanner crab, the morphol-
ogy and histology of the gastrointestinal tract is examined
and compared to that of the blue crab Callinectes sapidus
Rathbun. Three histological stains were used: hematoxylin
and eosin, periodic acid-Schiff (PAS), and the Feulgen
reaction with picro-methyl blue. The foregut, midgut, and
hindgut were examined. The fore- and hindguts are both of
ectodermal origin, and exhibit similar cuticular layers, epi-
thelial cells, and tegmental glands. The endodermally derived
midgut and caeca differ significantly from the fore- and
hindgut both in their lack of cuticle, and in the vacuolation
of the epithelial cell nuclei. One morphological difference
that was noted between the tanner and blue crabs was the
absence of aborizations in the posterior midgut caecum of
the tanner crab. The function of this caecum may be for
osmoregulation. Prolonged osmoregulation in brackish and
fresh water occurs to a significant extent in the blue crab
but not in the tanner crab because it remains in a marine

environment. This difference in habitats may explain the
variation in caecum structure. In most other aspects, the
histology and morphology of C. sapidus closely resembled
those of C. bairdi.

THE EFFECT OF POPULATION DENSITY ON THE GROWTH
OF THE BUTTER CLAM SAXWOMUS GIGANTUS

THOMAS C. KLINE

Division of Aquaculture and Invertebrate
Fisheries, School of Fisheries
University of Washington
Seattle. Washington 98195

Butter (or smooth Washington) clams, Sax idomus gigan-
teus (Deshayes), were grown for 2 yr at 4 population densi-
ties (96, 48, 24, and 12 clams/0.25 m 2 plots) in a Latin
Squares arrangement at the — 0.5-m tide level (MLLW) on a
privately owned beach approximately 1 km west of Port
Gamble on Hood Canal in Washington State. The clams,
dug up from within 10 m of the experimental site, and were
individually numbered and measured in length, width, and
thickness to the nearest 1 mm and placed into three groups,
each containing one third of the naturally occurring popula-
tion, depending on the clam length. The medium sized group
ranged from 76 to 80 mm, with the small and large groups
taking the remainder. The plots were filled by randomly

selecting from the three groups, with one third of each plot
represented by each of the three size groups. The clams were
planted in 1978 during the spring tidal series closest to
the summer soltice. They were removed, remeasured and
replanted at a similar tide in 1979. In 1980, the clams were
removed for the last time, during the soltice tidal series. In
order to compare the growth differences in the 4 population
densities, Walford plots of length at one time versus length
at another were made. Walford plots were also made for
width and for the product of length and width. The result-
ing plots showed that there was an appreciable difference in
growth between the 48 and 24 clams/plot. The 96 clams/
plot had the same growth slope as the 48/plot. The differ-
ence between the 12 and 24 clams/plot was also negligible.
The data indicated that the maximum density for best
growth is 24 clams/0.25 m 2 (96/m 2 ). The experiment
also demonstrated the usefulness of Walford plots to
optimize population in a grow-out situation as used in
shellfish aquaculture.

Olympia. Washington, September 10-12, 1982

Abstracts. 1982 NSA West Coast Section Meeting 1 1 3

THE FEEDING BEHAVIOR OF THE TEREBELLID

POLYCHAETE THELEPUS CRISPUS JOHNSON

IN RESPONSE TO CURRENTS

NANCY MUSGROVE

Division of Aquaculture and Invertebrate
Fisheries, School of Fisheries,
University of Washington
Seattle, Washington 98195

The role of currents in determining the feeding behavior
of Thelepus crispus was investigated as part of a large-scale
research project on the response of bottom-dwelling com-
munities to organic enrichment and pollution. Live worms
were collected from the intertidal beach at Garrison Bay on
San Juan Island, WA. They were placed in natural sediments,
in specially designed flow tanks at the Seattle Aquarium and
at the University of Washington Friday Harbor Labs. After
the worms reconstructed their tubes, the feeding behaviors
were observed under three different current velocities ranging
from 1 to 8 cm/sec. Particle settlement experiments were
also conducted at the three velocities to determine if flow
affected the settlement of food around the feeding worms.
To clarify any morphological limitations which might affect
the choice of food or feeding method in Thelepus, the
tentacles of preserved specimens were examined under a
scanning electron microscope. To corroborate findings in
laboratory experiments, field observations and How measure-
ments were made using SCUBA gear at Garrison Bay, WA.
When Thelepus is exposed to different current velocities it
orients its feeding tentacles in response to the direction of
flow and the areas of maximum particle settlements. At
speeds < 2 cm/sec, particle settlement is relatively even
around the worm mounds and Thelepus spreads it tentacles
in all directions on the sediment as well as in the water
column. It is under this type of flow condition that Thelepus
is abundant in the field. Suspension feeding may play an
important role in food gathering for Thelepus. At higher
current speeds (4 to 8 cm/sec) particle settlement becomes
differentiated between upstream and downstream areas
around the worm. The upstream face of the mound has
relatively few particles settling out. The downstream face
and area immediately behind the worm mound has greater
amounts of particles settling out. The placement of tentacles
mirrors the settlement patterns of particles. The strength of
the current is an important consideration as to how Thelepus
feeds and where it gathers its food.

VERTICAL MIGRATION OF GONYAULAX CATENELLA:
POTENTIAL IMPLICATIONS FOR MANAGEMENT OF
PARALYTIC SHELLFISH POISONING (PSP) PROBLEMS

LOUISA NISHITANI AND
KENNETH CHEW

School of Fisheries
University of Washington
Seattle. Washington 98195

The diel vertical migration pattern of the dinoflagellate

Gonyaulax catenella Whedon et Kofoid which produces

paralytic shellfish poisons, may have important implications
for management decisions by industry, public health agen-
cies, and research groups. This migration pattern influences
the length of time shellfish at different tide heights or
depths below rafts are exposed to G. catenella. The exposure
should be considered by health agencies, along with tide
height or depth, when planning routine sampling and by the
shellfish industry when selecting bivalve species to plant or
dredging depths. Because the vertical migration pattern is
greatly affected by the degree of stratification of the water
a predictive model which involves field studies of the effects
of changes in density gradients on density of G. catenella
should be developed. The vertical migration pattern appears
to be extremely important in the development of large
populations of G. catenella in certain sheltered bays, from
which significant numbers of G. catenella may then be
exported to waters outside the bay. An understanding of
the functioning of such bays may be useful in determining
timing and sites for monitoring and in selection of sites for
controlling G. catenella with the parasite, Amoebophrya
(if laboratory tests indicate such control would be safe,
desirable, and feasible).

ABALONE AND SCALLOP CULTURE IN PUGET SOUND

SCHARLEEN OLSEN

Washington Department of Fisheries,
Point Whitney Shellfish Laboratory,
Brinnon, Washington 98320

Three new species were cultured at Point Whitney Shell-
fish Laboratory during 1979-82; the native pinto (or
threaded) abalone Haliotis kamtschatkana Jonas, the red
abalone Haliotis rufescens Swainson, and the purple hinge
(or giant) rock scallop Hinnites multirugosus (Gale). A pilot
hatchery system was developed and various culture condi-
tions, methods, and temperatures were investigated. Growth
of the pinto abalone was followed over a period of 3 yr in
the hatchery. Comparisons of growth and survival rates
between juvenile pinto and red abalone were investigated
over a one-year period. The pinto growth rate was affected

by the type of culture container used and by the presence or
absence of light. At one year of age, pinto abalone shells

averaged 20 mm. At two years, mean shell length was 37 mm,
and the oldest year-class averaged 59 mm at three years of
age. Various scallop culture methods, feeding densities and
container configurations affected the scallop growth rates.
Salinity tolerance was studied and salinities < 23 ppt were
detrimental to normal growth and survival. Field plantings at
Lopez Island, Port Blakley, Willapa Bay, Manchester. Belling-
ham Bay, and Point Whitney were studied for growth and
survival of juvenile rock scallops. Growth rates of 4.2 mm/mo
were achieved at some locations.

1 14 Abstracts, 1 982 NSA West Coast Section Meeting

Olympia, Washington, September 10-12, 1982

PSP: ITS HISTORY, PROCESSES AND IMPACTS
AS APPLICABLE TO PUGET SOUND

TIMOTHY SAMPLE

METRO, Water Quality Division

Seattle, Washington 98104

This report provides a synopsis of available information
concerning the history, processes, and impacts associated
with paralytic shellfish poisoning (PSP) in Puget Sound.
Paralytic shellfish poisoning is a form of food poisoning in
which extremely lethal toxins, produced by certain dino-
flagellates, are accumulated in shellfish and passed on to
humans. Outbreaks of PSP appear to be spreading to previ-
ously unaffected areas. They are increasing in intensity
worldwide as well as within the Puget Sound basin. This
report includes a review of these trends and of the current
toxicity monitoring program established in the state of
Washington to protect the public from PSP. Attention is
also given to what causes toxic dinoflagellate blooms, partic-
ularly dinoflagellate cysts, and contributing environmental
factors (i.e.. temperature, precipitation, and nutrients).
Apparently, numerous environmental factors may influence
development of a bloom from newly emergent germlings.
In addition, the introduction of certain organic compounds,
called chelators, to coastal waters may create an environ-
ment favoring growth of the dinoflagellate population by
controlling the availability of certain growth-regulating trace
metals. A discussion of the nature of dinoflagellate toxins
and their possible effects on man and other organisms is
included. The recent discovery that dinoflagellate toxins
may be lethal to organisms other than man has serious impli-
cations: for example, consumption of toxic shellfish may
prove fatal to certain species of birds. Additionally, recent
investigations indicate that lethal levels of dinoflagellate
toxins can be accumulated, retained, and passed up the
food chain by herbivorous zooplankton that feed on toxic
dinoflagellates.

COMMERCIAL MARICULTURE OF A BAY SCALLOP

ARGOPECTEN CIRCULARIS (SOWERBY) IN THE

ENSENADA OF LA PAZ, BAJA CALIFORNIA

SUR, MEXICO

A. KIMBROUGH SIEWERS

Cultivos Marinos de Baja California
S. A. de C. V. RioNazas 163-401
Mexico 5, D. F. (and)
Pigeon Point Aquaculture Center
921 Pigeon Point Road
Pescadero, California 94060

Mexico's first private shellfish aquaculture company was
formed in La Paz, BCS. A local bay scallop, the Pacific calico
scallop Argopecten circularis, is grown in lantern nets

suspended from long lines. Scallop spat are collected by
putting sticks of plastic mesh in nylon "onion bags" which
are tied five to a weighted line and hung from long lines.
Collectors are set out in the spring and the seed scallops are
removed 2 to 5 mo later. Significant numbers of scallop
spat also regularly set on the lantern nets. Seed scallops are
grown in pearl nets during the nursery phase of culture, then
grown to market size in lantern nets. Fouling is removed
from the nets by a saltwater spray from a gasoline-powered
water pump. Scallops are stocked at a density of 25/0.1 m 2
(50 per level) for the final growth stage. Market size (5 to
6 cm) is reached in 5 to 7 mo. Four metric tons of scallops
were marketed in Mexico City in the first year of production.
A pufferfish, Spheroides annulatus (Jenyns), preyed on
cultured scallops by chewing open the bottom compartments
of some lantern nets. This was alleviated by shortening the
lantern nets by 3 levels. A hatchery was constructed, and in
the first experiment scallops were conditioned, spawned,
and the larvae reared to juvenile stage. Improvements in the
grow out system should include using 5-level lantern nets
in 2 mesh sizes (12 and 21 mm), and submerging the long
lines by 0.5 m. An annual production of 10 tons appears
necessary for profitability, with 20 to 30 tons possibly
optimum.

PSP RESEARCH: RECENT ADVANCES IN

ANALYTICAL AND BIOCHEMICAL

STUDIES

JOHN J. SULLIVAN AND
WAYNE T. IWAOKA

Institute for Food Science and

Technology
School of Fisheries
University of Washington
Seattle, Washington 98195

Paralytic shellfish poison (PSP), or "Red Tide," is a
persistent problem in the northern coastal areas of the
United States and monitoring of shellfish is accomplished
via mouse bioassays. We have developed an alternate analyti-
cal technique for measuring the toxins using high pressure
liquid chromatography. Comparison of both techniques
showed high correlation when toxin content in shellfish
samples contained about 60 /ig toxin per 100 g meat. The
mean variation was 25% when higher amounts of toxin
were present. Variation in the mouse bioassay is ± 20%.
Preliminary and proposed work will be reported on the
biochemical aspects of uptake, storage, and release of the
PSP toxins in shellfish.

Olympia, Washington, September 10-12, 1982

Abstracts, 1982 NSA West Coast Section Meeting

115

DISASTER AHEAD FOR THE YAQUINA
BAY OYSTER INDUSTRY?

LOUIS WACHSMUTH

Oregon Oyster Company
208 SW Ankeny Street
Portland, Oregon 9 7204

After 115 years of fishing and farming, the future of
Yaquina Bay is as uncertain and bleak as ever, with one
exception. The history of this bay, located in Newport, OR,
parallels histories of other west coast growing areas. The
oyster schooners from San Francisco, the old-time oyster
tongers, the replacement of the native Pacific oyster Ostrea
lurida Carpenter by the giant Pacific oyster Crassostrea gigas
(Thurnberg), the wood products pollution, the local town's
sewage, the infamous tidal wave, and the massive siltation
problem are all elements and events of the past 1 15 years.
The current crisis seems to be of major proportions and
threatens the future of oyster farming. Generally speaking,
oysters are no longer growing to full potential. Kumamoto
oysters (variants of C. gigas), which were grown on the

bottom 15 years ago, now grow only from rafts. Giant
Pacific oysters, as of 8 years ago, became stunted after
the second year of growth, only putting on thick layers
of blistered shells that were filled with a foul-smelling
exudate. They seldom reached "medium" size even after
6 years. Perhaps related to this is the fact that several other
forms of sea life have almost disappeared from our area
over the past 30 years. The source of this problem is
unknown, but could be related to the destruction of the
ocean food chain over the years. The stunting problem also
has been observed in other locations on the west coast. The
only ray of hope for this company is to repeat the great
switch of the 1920's. That is, change species of oysters once
again. The Japanese oyster, Crassostrea ariakensis (Wakiya)
(= Ostrea/ Crassostrea rivularis), seems to be the answer.
After experimenting for five years, we discovered these
advantages: (1) 50% faster growth than C. gigas, thereby
shortening the growth cycle by one year; (2) good flavor;
(3 ) absence of the stunting and blistering problem ; (4 ) larger
maximum size than C. gigas; (5) higher spawning tempera-
tures resulting in a firm and tasty meat during August and
September; and (6) uniform shell shape and attractive
interior shell surface.

INFORMATION FOR CONTRIBUTORS TO THE
JOURNAL OF SHELLFISH RESEARCH

Original papers dealing with all aspects of shellfish
research will be considered for publication. Manuscripts
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carefully prepared in the style followed in Volume 3,
Number 1, of the Journal of Shellfish Research (1983)
before submission to the Editor. Papers published or to
be published in other journals are not acceptable.

Title. Short Title, Key Words, and Abstract: The title
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JOURNAL OF SHELLFISH RESEARCH

Vol. 3, No. 1 June 1983

CONTENTS

Brian F. Beat

Predation of Juveniles of the Hard Clam Mercenaria mercenaira (Linne) by

the Snapping Shrimp A Ipheus heterochaelis Say and A Iphens normanni Kingsley 1

Rodney Dal ton and Winston Menzel

Seasonal Gonadal Development of Young Laboratory-Spawned Southern

(Mercenaria campeehiemis) and Northern (Mercenaria mercenaria) Quahogs

and their Reciprocal Hybrids in Northwest Florida . : 11

Paul J. Flagg and Robert E. Malouf

Experimental Plantings of Juveniles of the Hard Clam Mercenaria mercenaria (Linne)

in the Waters of Long Island, New York 19

/ D. Andrews

Transport of Bivalve Larvae in James River, Virginia 29

Catherine Enright, Donna Krailo, Larry Staples, Maria Smith, Carl I aughan, Debra Ward,
Pamela Gaul, and Elisabeth Borgese

Biological Control of Fouling Algae in Oyster Aquaculture 41

Mary L. Swift and Mohammed Ahmed

A Study of Glucose, Lowry-Positive Substances, and Triacylglycerol

Levels in the Hemolymph of Crassostrea virginica (Gmelin) 45

Edward R. Urban, Jr., Gary D. Pruder and Christopher J. Langdon

Effect of Ration on Growth and Growth Efficiency of Juveniles of

Crassostrea virginica (Gmelin) 51

Aurora Ledo, Enrique Gonzalez, Juan L. Barja and Alicia E. Toranzo

Effect of Depuration Systems on the Reduction of Bacteriological Indicators

in Cultured Mussels (Mytilus editlis Linnaeus) 59

C. B. Calloway and R. D. Turner

Documentation and Implications of Rapid Successive Gametogenic Cycles and

Broods in the Shipworm Lyrodus floridanus (Bartsch) (Bivalvia. Teredinidae) 65

RESEARCH NOTE

C. F. Phleger and S. C Cary

Settlement of Spat of the Purple-Hinge Rock Scallop Hinnites multirugosus (Gale)

on Artificial Collectors 71

Abstracts of Technical Papers Presented at the 1982 Annual Meeting National Shellfisheries

Association, Baltimore, Maryland - June 14-17, 1982 75

Abstracts of Technical Papers Presented at the 1982 Annual Meeting National Shellfisheries

Association, West Coast Section, Olympia, Washington - September 10- 1 2, 1982 105

COVER PHOTOMICROGRAPH: Female specimen of Alpheus heterochaelis Say (Decapoda; Alpheidae)
collected 26 June 1982 from an oyster reef near Beaufort, North Carolina, USA (scale bar = 5 mm). Photo-
graphed with 4 X 5-inch Graphic (Graflex) camera and Xenar lens (# 1 : 4.7/1.35) using Kodak Tech Pan
2415 film and processed in HC 110, F-dilution. (Exposure = 5 sec at f 45.) [Photograph provided by Henry E.
Page, University of North Carolina, Institute of Marine Science, Morehead City, North Carolina./

JOURNAL OF SHELLFISH RESEARCH

VOLUME 3, NUMBER 2

DECEMBER 1983

moratory

LIBRARY
AUG 26 1985

5 Hoie, Mass.

The JOURNAL OF SHELLFISH RESEARCH (formerly Proceedings of the

National Shellfisheries Association) is the official publication of the

National Shellfisheries Association

Editor-in-Chief

Dr. Roger Mann

College of William and Mary

Virginia Institute of Marine Science

Gloucester Point, Virginia 23062

Managing Editor

Dr. Edwin W. Cake, Jr.
Gulf Coast Research Laboratory
Ocean Springs, Mississippi 39564

National Shellfisheries Association
Publications Committee

Prof. Melbourne R. Carriker
College of Marine Studies
University of Delaware
Lewes, Delaware 19958

Dr. Rober E. Hillman

Battelle

New England Marine

Research Laboratory
Duxbury, Massachusetts 02332

Mr. Michael Castagna
College of William and Mary
Virginia Institute of Marine Science
Eastern Shore Laboratory
Wachapreague, Virginia 23480

Dr. Richard A. Lutz

Department of Oyster Culture

New Jersey Agricultural Experimental Station

Cook College, Rutgers University

New Brunswick, New Jersey 08903

Journal of Shellfish Research

Volume 3, Number 2
ISSN: 00775711
December 1983

Journal of Shellfish Research, Vol. 3, No. 2, 117-128, 1983.

SYMBIOTIC ASSOCIATIONS INVOLVING THE SOUTHERN OYSTER DRILL

THAIS HAEMASTOMA FLORIDANA (CONRAD) AND '" ^ f,> -

LIBRARY

MACROCRUSTACEANS IN MISSISSIPPI WATERS,

EDWIN W. CAKE, JR.

Oyster Biology Section

Gulf Coast Research Laboratory

Ocean Springs, Mississippi 39564

AUG 26 1985
.Woods Ho/* ka»*

ABSTRACT The symbiotic relationships between the southern oyster drill Thais haemastoma floridana (Conrad) and
two species of crabs, the blue crab Callineetes sapidus Rathbun and the striped hermit crab Clibanarius vittatus (Bosc),
were investigated in Mississippi. The crabs provided passive transport and food (attached fouling organisms) for the attached
drills; 99 blue crabs_ carried 203 drills (X= 2.0 ± 2.1 drills crab , range = l-17;mode = 1, N = 55 crabs); 233 hermit crabs
carried 299 drills (X= 1.3 ± 0.8 drills crab" 1 , range =1-6; mode = 1, N = 194 crabs). Drills attached to blue crabs were
twice the mean height and six times the mean weight of those attached to hermit crabs (36.8 mm and 8.9 g versus 18.5 mm
and 1.4 g. respectively). During one survey period 30 of 423 blue crabs (7.19"c) and 97 of 1,360 hermit crabs (7.17r) carried
drills. The oyster drill/blue crab symbiosis persisted while spawning female crabs congregated around Mississippi's offshore
barrier islands during the early fall of 1980 and ceased when the crabs died or migrated to deeper water during late fall.
The oyster drill/hermit crab symbiosis was continuous. Drills attached while the crabs were buried at the seawater/ substrate
interface, resting under peat outcroppings, or while scavenging among grass roots and jetsam. Once mounted, the drills
were not readily dislodged by movement of the crabs. In the laboratory drills more readily mounted hermit crabs with
attached drills and/or acorn barnacles than hermit crabs without these organisms. A typical mounting took only seconds to
complete; drills readily attached to moving hermit crabs. Drills dismounted from hermit crab shells when in the immediate
vicinity of live oysters. The drills preyed on acorn barnacles (Chelonihia patula [Ranzani] , Balanus spp.), oysters (Crasso-
strea virginica [Gmelin] , Ostrea equestris Say), and slipper shells (Crepidula spp.) that fouled the blue crab carapaces and
hermit crab shells. Two other gastropods (Cantharus cancellarius [Conrad] and Odostomia impressa [Say] ) were occasion-
ally attached to blue and hermit crabs that carried oyster drills.

KEY WORDS Blue crab, commensalism. macrocrustaceans. oyster drill, phoresis, striped hermit crab, symbiosis

INTRODUCTION

The southern oyster drill Thais haemastoma floridana
(Conrad) (Gastropoda; Muricidae) is the most destructive
predator of the American oyster Crassostrea virginica
(Gmelin) in coastal Gulf of Mexico waters from Florida to
Texas (Burkenroad 1931; St. Amant 1938; Butler 1953,
1954;Gunter 1953, 1979; McConnell 1953;Chapman 1955,
1958;Menzel and Hopkins 1955; Menzel et al. 1957, 1966;
Hofstetter 1959; May and Bland 1970; May 1971 ; Pollard
1973; Breithaupt and Dugas 1979). Mortalities of oysters
from drills can be as high as 50% a year (St. Amant 1938).
Drills prefer water salinities that usually exceed 18 to 20 ppt
(St. Amant 1938, Gunter 1979), and thus oyster reefs
located near open Gulf waters are subjected to drill preda-
tion during periods of drought or reduced freshwater inputs
(from extended closures of water impoundments). Offshore,
high salinity areas serve as reservoirs for drills; when inshore
salinities increase, the drills invade nearshore reefs as
planktonic veliger larvae. The larvae spend about a month
in the plankton and are widely dispersed (Butler 1953).
After metamorphosis, juvenile drills grow rapidly and can
grow an average of 28 mm a year (range = 20 to 42 mm)
(Butler 1953).

During the fall of 1980 I observed southern oyster drills
attached to many blue crabs (Callineetes sapidus Rathbun)
and gastropod shells occupied by striped hermit crabs

(Clibanarius vittatus [Bosc] ) in shallow waters around
Mississippi's barrier islands. The drills were being passively
transported by the crabs. St. Amant (1938) and Fothering-
ham (1976) reported this symbiotic relationship, but not to
the extent that 1 observed along Horn and Ship islands.
St. Amant found four and five drills attached to two blue
crabs in the vicinity of Grand Island, LA. He also noted
that drills occurred on flotsam . Fotheringham found juvenile
drills on 1 .7% of all gastropod shells (> 20 g) that were
occupied by C. vittatus along the Texas coast. Mark Chatry
(Louisiana Dept. Wildl. Fish., St. Amant Marine Laboratory,
Grand Isle, LA. pers. comm.) found drills on blue and striped
hermit crabs in lower Barataria Bay, LA, in the vicinity of
Grand Isle during 1980 and 1981. The late Capt. L. J.
Gorenflo of Biloxi. MS, photographed two blue crabs with
two and four drills attached, respectively, that were trawled
from Biloxi Bay channel in Mississippi Sound in 1953
(photograph provided by W. J. Demoran. Gulf Coast Research
Laboratory, Ocean Springs. MS). Capt. Gorenflo noted on
the photograph that most of the crabs were alive, but
some were weak and dead. His photograph is the only
evidence that this drill/crab symbiosis occurred previously
in Mississippi waters.

Other muricid oyster drills participate in similar drill/
crab symbioses along the Atlantic coast (Table 1). Federighi
(1931) reported that the Atlantic oyster drill Urosalpinx

117

118

Cake

cinerea Say utilized hermit crabs as a means of transport
in lower Chesapeake Bay. Harold Haskin reported (in
Carriker 1955) that on two occasions in Delaware Bay he
found five previously marked drills (U. cinerea) on shells
of the Atlantic moon snail Polinices duplicatus Say that
were inhabited by the flat-clawed hermit crab Pagimts
pollicaris Say. Some of the drills were attached to shells of
hermit crabs that were no larger than their own shells. One
marked drill attached to and was transported 3.5 in by a large
hermit crab within 15 minutes of release. Haskin suggested
that hermit crabs may play an important role in the distribu-
tion of oyster drills. MacKenzie (1962) reported that large
numbers of the thick-lipped oyster drill Eupleura caudata
(Say) and lesser numbers of U. cinerea were transported on
the carapaces of most horseshoe crabs (Limulus polyphemus
[Linnaeus] ) that he dredged from Long Island Sound. One
crab carried 761 thick-lipped and 4 Atlantic oyster drills. He
described their symbiotic association as phoresy. (Cheng
[1973] defined phorsey as a nonparasitic association in
which the smaller species, the phoront, is mechanically
carried on or in the larger species, the host, and no metabolic
interaction or dependency occurs.) Richards Nelson (in
Carriker 1955) reported as many as 140 Atlantic oyster drills
per horseshoe crab in New Haven Harbor, CT. Fred Sieling
(Maryland Dept. Nat. Resour., Annapolis. MD, pers.comm.)

and Michael Castagna (Virginia Inst. Mar. Sci., Wachapreague.
VA, pers.comm .(reported that they hadoccasionallyobserved
blue crabs transporting one or two thick-lipped drills in
lower Chincoteague Bay, VA, in the mid-1 950's. Federighi
(1931) suggested that oyster drills obtained food from
fouling organisms attached to the hermit crabs. Others
(St. Amant 1938, MacKenzie 1962) found no evidence of
drilling on the transport crab. Although several of these
authors alluded to a symbiotic association of macrocrusta-
ceans and muricid oyster drills, none attempted to docu-
ment or quantify the extent of those associations.

This paper describes the nature and extent of the drill/
crab symbioses that existed along Mississippi's barrier islands
during the fall of 1980. It examines the factors that initiate
and control these symbioses which appear to have character-
istics of commensalism and phoresis. Hereinafter, 1 shall
refer to the crabs as hosts and to the drills as symbionts.
Occassionally, I shall utilize the terms "infestation" and
"drill-infested" when presenting and discussing occurrence
data and when describing the existence of drills on the
shells of hosts. The use of these terms is not intended to
infer any parasitic relationship; they are simply utilized in
the absence of more appropriate terms. The crustacean and
molluscan taxonomies utilized herein follow those of
Williams (1965) and Abbott (1974), respectively.

TABLE 1.

A synoptic review of oyster drill/crab symbioses along the Atlantic and Gulf coasts of the United States.

Oyster Drill Species

Tliais haemastoma haysae
T. haemastoma

T. haemastoma

T. haemastoma floridana

Eupleura caudata

E. caudata
Urosalpinx cinerea
U. cinerea

U. cinerea
U. cinerea

Calotrophon ostrearum

Crab Species

Callinectes sapidus
C. sapidus

C. sapidus
Clibanarius vittatus
C. vittatus
C. sapidus
C. vittatus 2
Limulus polyphemus
C. sapidus

L. polyphemus
L. polyphemus
L. polyphemus

"Hermit crabs"
Pagurus pollicaris

Pagurus impressus

Locality

Grand Isle. LA

Mississippi Sound. Ocean Springs, MS

Lower Barataiia Bay. LA

Texas coast

Horn and East Ship islands, MS

Lower Chincoteague Bay, VA (1956)

Long Island Sound, NY
Long Island Sound, NY
New Haven Harbor, CT

Lower Chesapeake Bay, VA
Delaware Bay, DE

Reference

St. Joseph Bay, FL

St. Amant 1938

(L. J. Gorenflo, 1953

photograph)

Mark Chatry, LA DW&F,

pers. comm. 1981

Fotheringham 1976

(Present study)

Fred Sieling (MD DNR),
Mike Castagna (VIMS),
pers. comm. 1981
MacKenzie 1962
MacKenzie 1962
Richards Nelson (in
Carriker 1955)
Federighi 1931
Harold Haskin (in
Carriker 1955)
(E. W. Cake 1981, field
observation)

Identified as Tliais floridana haysae.
"Occupying the following gastropod shells: Busy con contrarium
alatus. and Tliais haemastoma floridana.
Occupying the shells of P. duplicatus.
Occupying the shells of 5. alatus.

B. spiratum plagosum. Murex fulvescens. Polinices duplicatus, Stromhus

Southern Oyster Drills Infest Macrocrustaceans

119

materials and methods

Blue crabs and striped hermit crabs with attached oyster
drills were collected at four stations on Horn and Ship
islands; those islands form the southern boundary of
Mississippi Sound (Figure 1). The crabs were collected in
shallow water (< 1 m) with dip nets or crab tongs. On
several occasions, all crabs encountered were collected to
determine the incidence of infestation. Field observations
were made on the behavior of the crabs and drills in their
shared habitats. The drills and potential prey items on the
crabs' shells (e.g.. acorn barnacles, oysters, and slipper shells)
were examined for evidence of predation. Infested crabs
and their attached drills were placed in individual plastic-
bags and transported alive in coolers to the Gulf Coast
Research Laboratory. Ocean Springs. MS. where they were
measured and weighed, and the numbers of drills, barnacles,
oysters, and slipper shells per crab (shell) were determined.

In the laboratory, studies of the oyster drill/hermit crab
symbiosis were conducted in 70- to 95-C all glass aquaria
using sand, seawater, and animals from Horn Island. The
experimental crabs occupied the shells of the lightning whelk
Busy con contrarium Conrad, the pear whelk B. spiratum
plagosum (Conrad), the southern oyster drill T. h. floridana,
and the Atlantic moon snail Polinices duplicates (Say), and
each was initially infested with acorn barnacles (Balanus spp.)

Each trial utilized 5 to 8 crabs. 40 to 50 drills (height
range. 15 to 75 mm), and lasted 2.0 to 3.5 hours. Various
combinations of crabs (with and without attached barnacles),
substrates (sand and oyster shells), oysters (live and empty
shells), and in-tank locations of same were utilized during
the experiments. Observations were made on the behavior
of the drills in relation to the crabs and oysters.

RESULTS

Description of Habitat

Independent drills and crabs and infested crabs shared
habitats in the inlets to Horn Island lagoons and in adjacent
shallow waters of Mississippi Sound (Figure 1). Those
habitats consisted of ( 1 ) exposed roots of salt-marsh grasses;
(2) submerged grassbeds and root-debris mats; (3) shallow
depressions in sandflats and under solid jetsam (e.g., boards
and timbers); (4) crevices in and ledges under peat out-
croppings; (5) small clumps of oysters; and (6) large groups
of oysters attached to submerged structures (e.g., wrecked
vessel debris, tree stumps, etc.). The drills and crabs fre-
quently made contact in those habitats, especially when
the drills crawled across partially buried blue crabs or
quiescent hermit crabs. Drills, crabs, and infested crabs
were also trawled-up together from Dog Keys Pass at the
west end of Horn Island ( Figure 1 ).

MISSISSIPPI SOUND

&&M ^

5 km

i I i I i_)

89°45' GULF OF MEXICO

Figure 1. Location of stations where drill-infested blue crabs and striped hermit crabs were collected.

120

Cake

Mean water salinities and temperatures in the study area
were 28.4 ppt (24.0 to 30.5 ppt) and 26.TC (20.0 to 28.0°C),
respectively, when drill-infested blue crabs were collected
(October 1980), and 21.5 ppt (20.0 to 30.5 ppt) and 19.4°C
(18.5 to 27.0°C), respectively, when drill-infested hermit
crabs were collected (October and November 1980).
Because of visibility and collecting device limitations, all
collections were made at depths of 1 m or less. The pre-
dominant substrate was well sorted and rounded quartz
beach sand, except in the lagoon inlets and along parts of
island shorelines where relict peat outcroppings existed.

Oyster Drill/Blue Crab Symbiosis

Ninety-nine infested blue crabs (98 9, 1 6) were collected
from four stations on Horn and East Ship islands on 2, 7, 9,
and 14 October 1980. All crabs were adults, while the
majority of the drills were juveniles. Mean sizes and weights
of the crabs and drills are given in Table 2. The crabs
carried a total of 203 drills (X = 2.0 ± 2.1 drills crab" 1 ,
range = 1-17) (Figure 2); the drills were attached to the
carapace (200), the chelae (2), and the abdomen (1). The
drill infestation mode was 1 drill crab -1 (N = 55 crabs;
55.5% of total); 22 crabs (22.2%) carried 2 drills apiece;
14 (14.1%) carried 3 drills apiece; 2 crabs each (2.2%)
carried 5, 6, and 7 drills apiece, respectively; and 1 crab
each (1.1%) carried 9 and 17 drills apiece, respectively
(Figure 2). No drill-infested blue crabs were observed
during three surveys in November (1.2, and 3 November
1980) and none was seen during numerous surveys during
the summer and fall of 1981.

On four occasions at two stations on Horn Island (Stn.
1.2 and 1.3, Figure 1) all blue crabs encountered were
collected. Seven percent (30 of 432) of the crabs carried
a total of 44 drills (Table 3).

Results of regression analyses of the drill infestation and
drill/crab meristic data are presented in Table 4. Only a
weak correlation existed between the number of attached
drills and the three crab meristics tested (carapace width,
weight, and the cross product of the width and weight).
In general, however, the larger the crab the larger the
number of attached drills.

Other Symbionts on Drill-Infested Blue Crabs

The most abundant epizoan on the drill-infested blue
crabs was the symbiotic acorn barnacle Chelonibia patula
(Ranzani) (see Overstreet 1978, 1982). Each crab carried a
mean of 81.8 ± 33.8 (12 to 287) live barnacles on its entire
exoskeleton and 35.2 ± 23.7 (2 to 122) live barnacles on
its carapace. The numbers of live barnacles on the entire
crab and also on the carapace alone were negatively corre-
lated with the number of attached drills (Table 4). Thus,
the larger the number of barnacles, the smaller the number
of attached drills (i.e.. barnacles reduce the space available
for attaching drills). Crabs with light barnacle infestations
carried 1.4 and 1.7 times as many drills as those with

moderate and heavy infestations, respectively; and crabs
with moderate barnacle infestations carried 1 .2 times as
many drills as those with heavy infestations (Table 5).
Thirteen (13.1%) of 99 drill-infested blue crabs had recently
dead (empty) barnacles (C. patula) on the carapace or
abdomen (X = 30 barnacles crab" 1 , range = 1 to 8, N =
39 barnacles). Two oyster drills were observed feeding on
barnacles attached to two crabs during the study, but the
barnacles did not appear to be an important food source for
the drills in general; only 39 of 8,099 (0.5%) barnacles on
the 99 drill-infested crabs were dead (empty).

Two (2.2%) of the 99 drill-infested crabs also carried one
specimen each of the buccinid gastropod Cantharus
cancellarius (Conrad), a common omnivore of mud/sand
bottoms in high salinity areas of Mississippi Sound. (Five

TABLE 2.

Summary of data from crabs that were infested with oyster drills
at four stations on Horn and East Ship islands, Mississippi.

Striped

Category

Blue Crabs 1

Striped Hermit Crabs 2

Total number crabs

423

1,360

Number infested crabs

Percent infested

7.10%

7.13%

Number drills

119

Mean number drills

on infested crabs

1.47

1.23

Mean number drills

on all crabs

0.10

0.09

'Combined data: Chimney Lagoon, Stn. 1.2 (7 & 14 October 1980)
and Ranger Lagoon, Stn. 1.3 (9 & 14 October 1980).
Combined data: Chimney Lagoon. Stn. 1.2 (3 November 1980)
and Ranger Lagoon. Stn. 1.3 (2 & 3 November 1980).

other blue crabs in addition to the 99 drill-infested crabs
were infested with specimens of C. cancelhrius only.)
Fourteen (1 4.1%) and three (3.3%) of the 99 drill-infestedcrabs
were also infested with single slipper shells (Crepidula spp.)
and pyram shells (OJostomia impressa [Say] ), respectively.

Oyster Drill/Hermit Crab Symbiosis

Two hundred thirty-three drill-infested striped hermit
crabs were collected at four stations on Horn and East Ship
islands on 2, 7. 9, and 14 October and 1, 2, and 3 November
1980 (Table 2. Figure 3). The hermit crabs occupied the
shells of 100 oyster drills (T. h. jloridana) (42.9%), 70
lightning whelks (B. contrariwri) (30.0%), 42 moon snails
(P. duphcatiis) (18.0%), 17 pear whelks (B. s. plagosum)
(7.3%), 2 giant eastern murexes (Murex fulvescens Sowerby)
(0.9%), and 2 Florida fighting conchs (Strombus alatus
Gmelin) (0.9%). The hermit crabs carried a total of 299
drills (X = 1 .3 + 0.8 drill shell" 1 , range = 1-6). The infesta-
tion mode was 1 drill crab" 1 (N = 194 crabs, 833% of total);
22 crabs (9.4%) carried 2 drills apiece; 12 crabs (5.2%)
carried 3 drills apiece; 3 crabs (1 .3%) carried 5 drills apiece;
and 1 crab each (0.4%) carried 4 and 6 drills apiece, respec-
tively (Figure 3). Mean sizes and weights of the crabs
(including the shell and attached epifauna but excluding
the drills) and the drills are given in Table 2. In general.
the larger the size of the hermit crab shell, the greater the
number of attached drills and the larger the size of the
attached drills.

On three occasions at two Horn Island locations (Sta.
1.2 and 1.3, Figure 1) all of the striped hermit crabs
encountered were collected. Seven percent (97 of 1,360) of

122

Cake

TABLE 4.

Results of regression analyses on data from drill-infested crabs collected at four stations on
Horn and East Ship islands, Mississippi

Host Crab Species

Correlations (versus number drills)*

r-Value

F-Value

Regression Equation

Callinectes sapidus

Clibanarius vittatus

Carapace width

Total crab weight

Cross product (width X weight)

Number live barnacles crab

Number live barnacles carapace

Maximum crab (shell) dimension
Weight of crab (+shell) versus

weight of individual drills
Cross product (size X weight)
Number live barnacles shell
Number live Crepidula shell

- 0.0346

0.166

Y =

2.8935 - O.0O55X

+ 0.0326

0.102

Y =

1.7755 +0.0018X

+ 0.0176

0.030

Y =

1.9339 +0.0050X

-0.1699

2.883**

Y =

2.6023 - 0.0067X

- 0.2508

6.508**

Y =

2.8419 - 0.0225X

+ 0.1836

8.062**

Y =

0.9212 +0.0044X

+ 0.2353

17.409**

Y =

0.5851 +0.0156X

+ 0.2297

12.871**

Y =

1.0820 +0.0426X

+ 0.0160

0.059

Y =

1.2797 +0.0009X

+ 0.1373

4.438**

Y =

1.2268 +0.0218X

*(Unless otherwise indicated.)
**(F-Value significant at the <X= 0.05 level.)

TABLE 5.

Summary of oyster drill and barnacle infestation data from 99 blue crabs collected at four stations on

Horn and East Ship islands, Mississippi

Mean Number

Number

Drills Crab" 1

Number Live

Barnacles Crab

Relative Intensity*

Barnacle-to-Drill

Blue Crabs

Oyster Drills

(Range)

Barnacles

(Range)

of Barnacles

Ratio

156

2.26

(1-

17)

4,265

61.81

(12-

-180)

Light

27.4

1.59

(1-

- 3)

3,197

118.40

(37-

-219)

Moderate

74.5

1.33

(1-

- 2)

637

212.33

(156-

-287)

Heavy

159.6

Totals/Means:

203

2.05

(1-

-17)

8,099

81.81

(12-

-287)

39.9

♦Light = <25% of carapace covered; moderate = 25 to 50% covered; heavy = >50% covered.

the crabs carried a total of 1 19 drills (Table 3). The 97 drill-
infested crabs occupied 44 shells of the oyster drill T. h.
floridana (45.4%), 26 shells of the lightning whelk B.
contrarium (26.8%), 20 shells of the moon snail P. duplicates
(20.6%), 6 shells of the pear whelk B. s. plagosum (6.2%),
and 1 shell of the fighting conch S. alatus (1 .0%).

Regression analyses were performed on three host
categories versus the number and/or weight of attached
drills (Table 4). All three correlations were weak but
positive. In general, the larger the occupied hermit crab
shell, the larger the number and size of the attached drills.

Several noteworthy differences existed between the two
drill/crab symbioses (Table 2). Drills that were attached to
blue crabs were twice the mean height as those on hermit
crabs (36.8 versus 18.5 mm) and consequently, six times
the mean weight (8.9 versus 1.4 g). Infested blue crabs
carried more drills than hermit crabs (X= 2.0 ± 2.1 versus
1.3 ± 0.8 drills crab" 1 , respectively). The maximum number
of drills carried by a blue crab (17) was 2.8 times the
maximum number carried by a hermit crab (6). Drill-
infested blue crabs carried approximately 9.4 times as

many live acorn barnacles as drill-infested hermit crabs
(81.8 versus 8.7 barnacles crab" 1 , respectively); however,
the number of drills on blue crabs was inversely related to
the number of barnacles, and the number of drills on
hermit crabs was directly related to the number of barnacles
(Table 4). Although no additional collections were made,
the drill/hermit crab symbiosis continued into the fall
of 1981, whereas the drill/blue crab symbiosis was not
observed when spawning ceased and the onset of colder
water temperatures caused blue crabs to migrate into
deeper water (late fall, 1980).

Other Symbionts on Drill- Infested Hermit Crabs

Acorn barnacles (896 of Balanus spp.) and slipper shells
(603 of Crepidula spp.) were the most abundant epifaunal
organisms on drill-infested striped hermit crabs (Table 6).
The mean numbers (and ranges) of barnacles and slipper
shells per hermit crab shell were 8.7 ± 18.8 (1 - 120) and
5.7 ± 5.7 (1 — 26), respectively. Weak but positive correla-
tions existed between the numbers of live barnacles and
slipper shells on the hermit crab shells and the number of

Southern Oyster Drills Infest Macrocrustaceans

123

Figure 3. Shell of lightning whelk (Busycon contrarium) inhabited by striped hermit crab (Clibanarius vittatus) and infested with Five
southern oyster drills (Thais haemastoma floridana) and three spotted slipper shells (Crepidula maculosa). Height and weight of whelk
shell (including attached fouling organisms, except drills): 132 mm and HOg, respectively. Mean drill height and weight, (ranges): 18.1 mm
(11.0 - 31.0) and 1.1 g (0.2 - 3.7), respectively. Infested crab was captured in the inlet of Ranger Lagoon (Stn. 1.3), Horn Island, MS,
on 2 November 1980.

TABLE 6.

Epifauna of gastropod shells occupied by drill-infested striped hermit crabs at four stations on
Horn and East Ship islands, Mississippi

Mean and

Shell Species Occupied

(N)

Crepidula spp.

Range

Balanus spp.

Range

Ostrea equestris

Range

Busycon contrarium

(70)

385

5.5, 1-26

204

2.9, 1- 55

0.1,(1)

B. spiratum plagosum

(17)

108

6.4, 2-20

1.7. 1- 12

0.5, 1-7

Murex fulvescens

(2)

0.5, 1

1.5,(3)

Polinices duplicatus

(42)

1.3,1- 7

1.8, 1- 20

Strombus alatus

(2)

6.0,(6)

Tfwis haemastoma floridana

(100)
(233)

41
603

0.4, 1- 5
2.6, 1-26

587
896

5.9, 1-120

18
33*

0.2, 1-3

Grand totals, means, ranges

3.8, 1-120

0.1. 1-7

*9 live; 4 dead with right valve drilled; 20 dead with only left valve remaining.

attached drills (Table 4). In general, the greater the number
of slipper shells on a drill-infested hermit crab shell, the larger
the number of drills (Table 4). Thus, the presence of attached
prey species is directly related to the attractiveness of the
crab's shell to foraging drills. The positive correlation in the
case of barnacles on hermit crab shells (as opposed to the
negative correlation in the case of barnacles on blue crab
carapaces) is a function of the total shell space available for
foraging drills to attach. (Blue crabs heavily infested with C.
patula have limited space on the carapace for drills to attach.)
Several oyster drills were observed feeding on epifauna
attached to hermit crab shells. One 34-mm drill had rasped

a hole and was feeding on a 32-mm oyster spat (C. virginica)
which was attached to the outside of a 107-mm lightning
whelk shell when the host hermit crab was collected.
Another 36-mm drill was rasping a hole along the margin of
a 29-mm slipper shell {Crepidula plana Say) which was
attached inside the aperature of a 122-mm lightning whelk
shell when the host hermit crab was collected. Only 9
(27.3%) of 33 crested oysters {Ostrea equestris Say) found
on drill-infested shells occupied by hermit crabs were alive;
4 shells were empty and drilled by a muricid gastropod
(probably T. h. floridana); and 20 were represented only by
their attached left valves.

124

Cake

Additional Drill/Crab Symbioses

During the study, several additional examples of oyster
drill/crab symbioses were observed in the vicinity of Horn
Island, MS. Several large horseshoe crabs (L. polyphemus)
along the island's beach had one or two moderate-to-large
oyster drills attached to their abdomens. Two additional
drill-infested crab species were collected in commercial
blue crab traps in deeper water (> 3 m) off the island's
north beach. One stone crab {Menippe merceiiaria [Say] ;
95 mm, 184 g) carried five drills (66 to 71 m) and five live
barnacles (C. patula) on its carapace. One spider crab
(Libinia dubia H. Milne Edwards; 73 mm. 148 g) carried
one drill (62 mm, 34 g) and 85 live and 10 dead barnacles
(C. patula) on its carapace. Those symbiotic associations
may have been artificially produced, however, because the
crabs were confined in a trap that attracted and permitted
the entry of large numbers of oyster drills.

An additional oyster drill/hermit crab symbiosis was
observed during the summer of 1981 in St. Joseph Bay, FL.
Four red hermit crabs (Pagurus impressus [Benedict] )
occupying shells of the Florida fighting conch {S. alatus)
carried six mauve-mouth oyster drills (Calotrophon ostrearum

[Conrad] ) (Figure 4). The mean height of the conch shells
was 86 mm (78 — 94 mm) and the mean weight of the shell
plus crab was 76 g (52 - 93 g). The mean height and weight
of the drills were 21.0 mm (17.5 - 23.6 mm) and 1.0 g
(0.5 - 1.6 g), respectively. The conch shells were also
occupied by five crested oysters (O. equestris), one of
which was incompletely drilled, and numerous slipper shells
(Crepidula maculosa Conrad and C. plana) of various sizes.

Behavior of Oyster Drills and Hermit Crabs

When given the opportunity to interact with hermit
crabs in laboratory aquaria, the oyster drills behaved as
follows:

1. The drills more frequently mounted hermit crab
shells that had live barnacles attached, and also those that
had other drills attached. When live barnacles were present.
179 (91.8%) of 195 drills mounted hermit crab shells;
124 drills (63.6%) attached if other drills were already
attached to the hermit crab shells; and 55 drills (28.2%)
attached when no other drills were present. When no live
barnacles were present on the hermit crab shells, 1 1 drills
(5.6%) attached in the presence of other drills and 5 drills
(2.6%) attached in the absence of other drills.

Figure 4. Shell of fighting conch (Strombus alatus) inhabited by red hermit crab (Pagurus impressus) and infested with two mauve-mouth
oyster drills (Calotrophon ostrearum) and one spotted slipper shell (Crepidula maculosa). Height and weight of conch shell (including fouling
organisms, except drills): 94 mm and 93 g, respectively. Drill heights and weights: 21.4 and 23.6 mm, 1.2 and 1.6 g, respectively. Slipper
shell length and weight: 27.0 mm and 2.0 g, respectively. Infested crab was captured in the vicinity of Presnell's Fish Camp, St. Joseph Bay,
Port St. Joe, FL, on 15 June 1981.

Southern Oyster Drills Infest Macrocrustaceans

125

2. The usual drill-to-crab mounting occurred in the
following manner: When the hermit crab shell was
encountered, the drill raised its tentacles and siphon,
extended them forward, and examined the shell; the drill
then raised the forward portion of its foot, attached to the
shell, and when most of the foot was connected, it pulled
its body and shell onto the host's shell. Once mounted, the
drill usually moved around the shell for a few minutes
before becoming quiescent. The drill-to-shell mounts were
relatively fast and were completed approximately 5 seconds
after initial contact.

3. Most drills mounted the part of the hermit crab's
shell that was initially encountered, regardless of the position
and activity of the host crab's tentacles, eyes, and chelipeds.
Drills were able to mount hermit crab shells that were
moving when encountered.

4. Drills mounted hermit crab shells from sand and
solid substrates with relative ease; 61% of the mountings
were from sand and 39% were from aquarium sides, other
crab shells, dead oyster shells, and pieces of brick. Drills
also attached to passing crab shells while upside-down
(shell aperature up) in the sand.

5. On three occasions 15 drills mounted one hermit
crab shell (6 drills per hermit crab shell was the greatest
infestation observed in the field). Fifteen drills mounted one
crab within 50 minutes (0.3 drill min" 1 ). The greatest attach-
ment rate on one hermit crab shell was 1 1 drills within
6 minutes ( 1 .8 drills min" 1 ).

6. Apparently, oyster drills were attracted to barnacles
on the hermit crab shells and remained on the shell until
more preferred prey such as oysters were encountered
or until dislodged for other reasons. The drills dismounted
from hermit crab shells onto or immediately adjacent to
live oysters, but rarely remounted the crab shells once on
live oysters. Twenty-four (57.1%) of 42 drills in three
experiments were transported to live oysters by hermit
crabs.

DISCUSSION

Factors Controlling Oyster Drill /Crab Symbioses

Southern oyster drills were attracted to and mounted
blue crabs and striped hermit crabs for at least one of the
following reasons:

1 . Foraging and the presence of potential food.

The presence or probable presence of acceptable
prey species of the southern oyster drill appeared to be the
most important controlling factorin the drill/crab symbioses.
St. Amant (1938) and Butler (1953, 1954) reported that
drills, especially young ones, will consume barnacles, and
that drills of all sizes will prey on oysters and mussels.
During this study I observed direct and indirect evidence
of drill predation on epifauna of blue crab and hermit
crab shells. Direct evidence included actual feeding of drills

on barnacles (on blue crabs) and indirect evidence included
Thais drill holes in dead oysters and in-progress drilling of a
slipper shell (on hermit crab). This is the first known
evidence of slipper shell predation by the southern oyster
drill. All drill-infested blue crabs had live barnacles attached
to their exoskeletons, but if the crab's carapace was heavily
infested (> 50% coverage) with barnacles, space availability
appeared to limit the number of attached drills. The numbers
of barnacles and slipper shells on drill-infested hermit crabs
were, however, positively correlated with the number of
attached drills.

Foraging drills are negatively geotactic; they will
move upward when placed under water, unless they
encounter acceptable food in which case they remain with
the food species "indefinitely" (Butler 1979). The act of
crawling up onto any solid substrate including crab shells
or aquarium walls is a normal foraging behavior of oyster
drills. Butler (1979) reported that the South Australian drill
Thais orbita (Gmelin) moved up the walls of a container in
the absence of barnacles, but remained with and fed on
barnacles (Balanus glandula Darwin) when present. Whether
the drill's negative geotaxis was automatic or in response
to the release of metabolites by potential prey species was
not determined and, in the case of relatively small substrates
like crab shells, the two behaviors may be inseparable. In
the case of these drill/crab symbioses, most initial attach-
ments probably resulted from foraging, but were enhanced
if acceptable prey species were present.

2. The presence of other attached drills (gregarious
factor).

Southern oyster drills are normally gregarious,
especially during feeding and spawning when food by-
products and pheromones, respectively, are released (St.
Amant 1938, Gunter 1979). The presence of 16 drills on
the carapace of one blue crab is an example of gregarious-
ness (Figure 2). The 16 drills were clumped together;
however, only five live barnacles were present and no feeding
or spawning activities were in progress. Thus, some other
factor attracted and held the drills on the crab's carapace.
During the laboratory experiments, only 11 (5.6%) of 195
drills attached to crabs which had other attached drills
but no live prey (barnacles). Thus, this appears to be a
minor factor. When the initial field collections were made,
the drill spawning season had ended and no reproductive
activities were observed among the young drills used in
the laboratory behavior trials.

3 . The availability of solid, stable substrates for pro-
tection or shelter.

Oyster drills, especially recently settled juveniles,
are normally associated with and attached to firm substrates
such as oyster shells, rocks, and submerged objects (timbers,
stumps, etc.) for food (epifauna), protection (from
predators), and shelter (from currents, waves, and potential

126

Cake

loss of attachment and subsequent abrasion, burial, or
predation). Because of the dearth of such substrates in the
vicinity of the barrier islands, the attachment of young
drills to the crab shells may have been a defensive as well as
a foraging behavior. Small drills which were attached to
crab shells had a lower probability of being consumed
by fish and crab predators than unattached drills. Although
striped hermit crabs will kill oyster drills (Gunter 1979),
they are unlikely to leave the protection of their gastropod
shell to attack attached drills; however, small drills within
the aperature of the hermit crab shell may be subjected to
such predation. Blue crabs will remove attached drills if
they are within reach of the chelae and the crabs can dis-
lodge attached drills by "rubbing" them against aquarium
walls. In either case, protection is lost, and the drills may be
subject to predation.

4. The presence of eggs on ovigerous blue crabs.

Eggs or their by-products which are released from
ovigerous crabs may biochemically attract foraging drills.
Sixty-four (65.3%) of the 98 drill-infested females were
ovigerous, 26 (26.5%) were "spent" (the zoeae had recently
hatched), and the remaining 8 (8.2%) had not yet spawned.
The probability of drill infestation is greater when the
females are ovigerous than when they are not. Of 55 females
infested with a single drill. 31 (56.4%) were ovigerous;
16 (72.7%) of 22 females with two drills were ovigerous;
11 (78.6%) of 14 females with three drills were ovigerous;
and 6 (75.0%) of 8 females with five or more drills were
ovigerous. In a related study of drill damage to blue crabs
in commercial traps north of Horn Island. I observed several
drills feeding on the "sponge" of ovigerous females. The
highly protrusile proboscis of oyster drills permits them to
rasp and feed on crab eggs while attached to the carapace
and abdomen of ovigerous females.

5. The presence of biochemical stimulants or by-
products from wounded or moribund blue crabs.

Wounded, moribund, or dead crabs, especially blue
crabs, may represent a potential food source for the other-
wise carnivorous drills. On several occasions in November
1980, when large numbers of spawned-out females were
dead or dying, a few were stranded on the beach at low tide
with drills still attached to their carapaces. Were the drills
waiting for passive transportation to continue or were
they waiting for a meal? During a related study of drill
damage to commercially trapped blue crabs north of Horn
Island in the spring of 1981. I observed that drills pene-
trated the crabs' exoskeletons via wound holes, autotomized
appendage stumps, thin appendage joints, and occasionally
via holes drilled in the carapace. The drills also used their
protrusile proboscis to penetrate the thin membranes at
the bases of the gills within the branchial chambers to
gam access to thoracic muscle tissues. No such crab

predation was observed during the present study of 203
drills that were attached to 99 live blue crabs.

6. Increased random attachment to available substrates
by an exploding drill population.

Environmental conditions near the barrier islands
may have promoted the drill/crab symbioses. Extended
drought conditions during 1979-1981 increased salinities
in Mississippi Sound and permitted the settlement of
relatively large numbers of young drills in normally marginal
habitats. Those drills became abundant in habitats con-
taining barnacles, mussels, and oysters around the barrier
islands. The sheer abundance of the drills and their random
attachment to all firm objects may account for their
presence on crabs. In those instances when infestation
prevalence was determined, blue crabs and hermit crabs
exhibited the same prevalence (7.17c). Although I made no
attempt to document the presence of drills on flotsam and
jetsam around the barrier islands, Federighi (1931) and
L. A. Stauber (in Carriker 1955) reported that young oyster
drills (U. cinerea) were distributed by attaching to floating
algae as well as to other flotsam and jetsam. I routinely
observed drills on submerged planks and other discarded
items in barrier island lagoons during this study.

7 . In response to a programmed symbiotic phenomenon.

If the drill/crab symbioses are as well established as
shown by this and other studies (Table 1). then muricid
drills may be programmed to seek crabs for their trans-
portation potential. The availability of transportation to
unpopulated areas, especially those with abundant food
supplies, may foster the symbiotic relationship.

Probable Role of Macrocrustaceans in the Migration of Southern
Oyster Drills

Along the Gulf of Mexico coast, blue crabs and striped
hermit crabs are common inhabitants of estuaries and
oyster reefs (McDonald 1940, Galtsoff 1964, McClellan
1965, Fotheringham 1976, Bahr and Lanier 1981) where
they tolerate a wide range of water salinities and tempera-
tures (Christmas and Langley 1973). Blue crabs move
about extensively (Darnell 1959) and can travel as much as
1.6 to 2.0 km day" 1 (H. Perry, Gulf Coast Research Labora-
tory, Ocean Springs, MS, and M. Oesterling, Virginia
Institute of Marine Science, Gloucester Point, VA, unpub-
lished data). Thus, they could carry oyster drills from barrier
island habitats to inshore oyster reefs within a week. In
contrast, striped hermit crabs travel much less and usually
remain within the littoral and shallow, sublittoral zones
(Fotheringham 1975). They travel as much as 156 m day" 1
(Hazlett 1981) and. thus, could carry oyster drills (to
nearby oyster reef) but not as far as blue crabs. On the
other hand, oyster drills do not migrate (Butler 1953);
unless carried by crabs or other means, the drills probably
remain within the vicinity where they originally settled.

Southern Oyster Drills Infest Macrocrustaceans

127

In Mississippi Sound, at least three species of crabs
(blue, striped hermit, and horseshoe) were observed trans-
porting drills during this study. Thus, the drill/crab
symbioses may be important in distributing juvenile and
young adult drills. The quantity of drills transported by
this means is small when compared with the number of
larval drills that are distributed in the plankton to high
salinity areas following reproduction. Nevertheless, the
crabs might carry drills to areas where currents do not
carry larval drills, and they can transport drills throughout
the year.

Along the Atlantic and Gulf coasts of the United States,
at least four species of muricid oyster drills [Calotrophon
ostreanim, Eupleura caudata, Thais haemastoma floridana,
and Urosalpinx cinerea) and five species of arthropods
( Callinectes sapidus, Clibanarius vittatus, Pagiims impressus,
Pagitrus pollicaris, and Limulus polyphemus) (Table 1)
participate in drill/crab symbioses. Although relatively few
reports about these symbioses appear in the literature. I
suspect that they are common and have an important
role in extending the distributions of oyster drills.
MacKenzie (1962) concluded that horseshoe crabs (L.
polyphemus) were important distributors of Atlantic coast
oyster drills (E. caudata and U. cinerea) throughout Long
Island Sound and perhaps beyond. Harold Haskin (in
Carriker 1955) concluded that hermit crabs (P. pollicaris)
played an important role in the distribution and migration
of Atlantic oyster drills (U. cinerea) in Delaware Bay.

The distributory effects of these drill/crab symbioses
may be somewhat negated, however, because blue crabs
and striped hermit crabs prey on small oyster drills. Blue
crabs in Horn Island lagoons (pers. observ.) and in nearby
Lake Pontchartrain, LA (Darnell 1958), readily consume
small gastropods which they ingest whole. Gunter (1979)
reported that striped hermit crabs killed southern oyster
drills in Apalachicola Bay, FL, by pinching their tentacles
until they bled to death; thereafter, the crabs pulled the
drills from their shells, consumed the flesh and occupied
the newly emptied shell. Of 1,360 striped hermit crabs
collected during November 1980, from the Horn Island
lagoons (Stn. 1.2 and 1.3), 825 (60.7%) occupied shells
of the southern oyster drill. (The next most frequently
occupied shell was that of the moon snail P. duplicatus
[23.0%].) Rudloe (1971) documented the attack of a
striped hermit crab on a live pear whelk Busycon spiratum
(Lamarck) in which the crab killed the whelk with its
chelae, extracted and consumed the flesh, and occupied
the new shell briefly before returning to its "old" shell.

Drill/Crab Symbioses: Commensalism or Phoresis?

Cheng (1967) discussed the importance of commensalism
and phoresis in the marine environment and pointed out
that the two symbioses differed primarily with regard to
nutritional aspects. He defined "commensalism" as a more
or less intimate relationship in which the commensal (in

this case the drill) generally derives physical shelter from
the host (the crab), is nourished on food organisms that are
associated with but not a part of the host (barnacles, oysters,
slipper shells), and is not metabolically dependent on the
host. Literally, commensalism means "eating at the same
table." It is a loose type of nonobligatory relationship
(Cheng 1967). He defined "phoresis" as a loose, nonobliga-
tory relationship in which one species, the host (crab),
merely provides shelter, support, or transport for the other
species, the phoront (drill). Metabolic dependency is not
involved. In a more restrictive definition, Cheng (1973)
considered phoresis as an association in which the smaller
of the two species, the phoront. is mechanically carried in
or on the larger species, the host, and no metabolic inter-
action or dependency occurs. It does not involve a sharing
of food as does commensalism. According to Cheng's
definitions of phoresis, those animals, commonly referred
to as being epizootic or epizoic. are engaged in phoretic
associations with their hosts.

The symbiotic relationships between southern oyster
drills and crabs in Mississippi Sound share components of
commensalism and phoresis. The two symbioses can overlap
according to Cheng (1967). and this is apparently the case
with the drill/crab associations described herein. In a
limited sense, the drills derive passive transport (cf., phoresis),
shelter (cf., phoresis and commensalism), albeit negligible,
and support (cf., phoresis) from the crab hosts. The drills
derive nutritional benefit in a nonobligatory fashion (cf.,
commensalism) from the epifauna on the crab hosts, but
the drills do not "share" those prey species in the traditional
sense (cf.. commensalism) such as do hermit crabs and
attached sea anemones. On the other hand, if drills consume
eggs from ovigerous blue crab females or attack and kill
free-living blue crabs, then the relationship can be considered
predatory.

If food availability and utilization are the primary
controlling factors in the drill/crab symbioses, the relation-
ships should be categorized as modified forms of commens-
alism. On the other hand, if, as MacKenzie (1962) observed,
the drills primarily derive passive transport from the crabs,
the relationships should be categorized as modified forms
of phoresis. Cheng (1967) noted a considerable overlapping
between commensalism and phoresis, yet he provided no
examples of symbioses that shared characteristics of both.
He suggested that one type of symbiosis may evolve into
another. In that case, neither the commensalitic nor phoretic
behavior of the two symbionts appears to be dominating.
I suggest, therefore, that the commensalistic components
probably evolved first and the phoretic components occurred
secondarily. The drill/crab symbioses in Mississippi waters
appear to be primarily commensalistic and secondarily
phoretic, and perhaps should be defined as phoretic
commensalism. Of the seven controlling factors discussed
at the beginning of this section, foraging and the presence
of attached prey species (on blue and hermit crab shells)

128

Cake

and egg masses (on ovigerous blue crabs) probably initiated
the relationships', food availability, gregariousness, and sub-
strate stability (protection and/or shelter) probably pro-
longed them; and the foraging for new food sources or
dislodgment probably terminated the relationships. The
drill's "predatory" behavior toward wounded or moribund
blue crabs appeared to be an expression of the drill's
normal opportunistic feeding, especially when it occurred
in commercial crab traps. The possibilities of random
attachment to solid substrates and "programmed" trans-
portation attempts appeared to be the least plausible
controlling factors.

ACKNOWLEDGMENTS

I gratefully acknowledge the financial and logistical
support of the Gulf Coast Research Laboratory and the
cooperation and assistance provided by officials of the Gulf
Islands National Seashore (National Park Service). Roger
Jennings and Rick Sherrard assisted with field collections
and laboratory measurements; Gary Licht conducted prelim-
inary studies of oyster drills and hermit crabs: and Vincent
Smith provided occasional boat transportation and infested
specimens from his commercial crab traps. Valarie Hebert
provided statistical and computer assistance and Lucia
O'Tooleand Cindy Dickens typed the manuscript drafts.

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Journal of Shellfish Research, Vol. 3, No. 2, 129-134, 1983.

PREDATION ON AMERICAN OYSTERS (CRASSOSTREA VIRGINICA [GMELIN] )

BY AMERICAN LOBSTERS (HOMARUS AMERICAN VS MILNE-EDWARDS),

ROCK CRABS (CANCER IRRORATUS SAY), AND

MUD CRABS (NEOPANOPE SAYI [SMITH] )

ROBERT W. ELNER 1 AND RENE E. LAVOIE 2

1 Department of Fisheries and Oceans
Invertebrates and Marine Plants Division
Biological Station

St. Andrews, New Brunswick, Canada EOG 2X0

2 Department of Fisheries and Oceans
Fisheries Research Branch

Halifax, Nova Scotia. Canada B3J 2S 7

ABSTRACT Predation on the American oyster Crassostrea virginica (Gmelin) by the American lobster Homarus
americanus Milne-Edwards, the rock crab Cancer irroratus Say, and the mud crab Neopanope sayi (Smith) was studied in the
laboratory. When provided with a range of oysters from 10 to 35 mm shell length (SL), lobsters (55-98 mm carapace
length [CL)) and rock crabs (32-107 mm carapace width [CW]) could all open oysters of 25 to 30 mm SL, but they
usually selected smaller oysters. Oysters of 30 to 35 mm SL appeared to be a critical size as they were rarely preyed upon
by either lobsters or rock crabs. Although all lobsters had a similar broad preference for oysters of 10 to 25 mm SL, larger
rock crabs preferred larger oysters than smaller rock crabs. Predation rates were variable among individuals but were
generally faster in larger than in smaller lobsters and rock crabs. Maximum mean lobster and rock crab predation rates were
28.0 and 4.5 oysters • predator ■ day , respectively. Groups of rock crabs (32-58 mm CW) and mud crabs ( 14-23 mm
CW) that fed on oysters of 2 to 9 mm SL which were attached to spat collectors, averaged 0.59 and 0.44 oyster • crab -1 ■
day , respectively. Larger rock crabs (50-76 mm CW). foraging on spat collectors, consumed 1.26 oysters * crab
day . No discernible sexual differences existed in either oyster size selection or predation rates for the lobsters and crabs.
Patterns of oyster destruction were predator-specific. Lobsters opened oysters by indiscriminate crushing, whereas rock
crabs exploited weak spots around the shell margin and thin areas in the cup valve. Isolating oyster < 30 to 35 mm SL
from decapod predators and barring rock crabs and mud crabs from spat collectors would reduce oyster mortality.

KEY WORDS: Oysters, Crassostrea virginica, lobsters, crabs, predation. selectivity, mariculture

INTRODUCTION did not consider opening techniques or size-specific

predation rates.
The American oyster Crassostrea virginica (Gmelin) is In Caraquet Bay, NB, culturists collect oyster spat on

cultivated extensively on the eastern coast of North America "chinese-hat" collectors, so-called because of the conical
and has economic importance in many communities. Studies collector plates, the 0.33-m diameter plates are stacked in
of oysters have identified cancrid. portunid, and xanthid bundles of 12 with a gap of 30 mm between adjacent
crabs (Menzel and Hopkins 1955, McDermott 1960. Krantz P^tes. The bundles are suspended, from rafts or fences,
and Chamberlin 1978, MacKenzie 1981). oyster drills 0.3 m above the sea bottom. Culturists detach oyster spat
(Galtsoff 1964), and starfish (Galtsoff 1964) as the principle from the collectors at a shell length of about 23-25 mm
predators. (See MacKenzte [1981] for a comprehensive and use various P rotective reari "g techniques to grow the

oysters to seed. The oysters are exposed to lobsters and

review of oyster mortality factors.)

We found in our laboratory study that American lobsters

rock crabs when relaid as seed onto growing areas at shell

lengths from 25 to 60 mm.

(Homarus americanus Mhne-Edwards) are also predators of We conducted a laboratory investigation on the shell-

oysters. Previously it was shown that lobsters prey on op ening behavior and feeding rates of lobsters and rock
several types of molluscs and other invertebrates (Ennis crabs on unattached oysters from Caraquet Bay. Predation
1973, Elner and Jamieson 1979, Elner 1980, Scarratt 1980). by rock crabs and mud crabs {Neopanope sayi [Smith] ) on
The rock crab Cancer irroratus Say preys on oysters in oysters attached to chinese-hat collectors was also
Caraquet Bay, New Brunswick, Canada (R. W. Elner, considered. Both crabs are abundant on the collectors
unpublished data), and on other molluscs and invertebrates suspended in the bay and cause oyster mortality. We were
in other areas (Scarratt and Lowe 1972, Elner and Jamieson particularly interested in obtaining information about the
1979, Elner 1980). The only previous quantitative investi- predators that could be used to improve culture strategy
gation of oyster predation by rock crabs (MacKenzie 1981 ) and ultimately increase oyster yields.

129

130

Elner and Lavoie

MATERIALS AND METHODS

Lobsters and large rock crabs were captured by otter
trawl in Passamaquoddy Bay, New Brunswick, Canada, near
the mouth of the Bay of Fundy . Small rock crabs, mud crabs,
and oysters were collected from commercial oyster beds in
Caraquet Bay, NB. The oysters had been grown on the
cement substrate which coats the chinese-hat collectors,
and thus had cement cultch bonded in their left (cupped)
shell valves. During experiments, the predators and oysters
were kept in 0.35- X 0.5-m glass aquaria filled to a depth of
0.25 m with running seawater. The seawater temperature
was 13 ± 1°C and the salinity range was 29 to 32 ppt
throughout our investigations.

Lobster size was measured as carapace length (CL) from
the posterior edge of an eye socket to the posterior edge of
the carapace, parallel to the longitudinal axis. The sizes of
rock crabs and mud crabs were determined by measuring
carapace width (CW) between the tips of the distal marginal
teeth. Maximum shell dimension ("length") (SL) was
measured to express oyster size. All measurements are
accurate to the nearest 0.1 mm.

Predators were held without food for 2 days before
feeding experiments to standardize hunger levels. Only
uninjured, apparently healthy predators and oysters were
used.

Predation techniques used by lobsters from 55 to 98 mm
CL and rock crabs from 32 to 107 mm CW on oysters of
5 to 35 mm SL were observed. We also observed the tech-
niques used by small rock crabs (32—58 mm CW) and mud
crabs (14-23 mm CW) as they fed on oyster spat (2-9 mm
SL) which were attached to the chinese-hat spat collectors.
Shell fragments were collected to help interpret and describe
breaking techniques.

Individual lobsters and rock crabs from three and four
size groups, respectively , each group comprising six predators,
were presented with five size classes of oysters of ten individ-
uals each. The oysters were spread over the bottom of the
aquaria. Prey and predators sizes were:

Oysters (mm, SL): 10-15, 15-20, 20-25, 25-30, 30-35.
Lobsters (mm, CL): 55-63 (9), 58-62 (d), 85-98 (9).
Rock crabs (mm, CW): 32-46 (9), 35-45 (d), 73-79 (d),
94-107 (d).

The sizes of the predators were within the size ranges
that occur on oyster beds in Caraquet Bay. The predators
were segregated by sex to determine whether feeding
behavior or rates were different. The number of oysters
eaten in each size class was monitored daily for 11 days;
all oysters eaten were replaced by live oysters of the same
size class to maintain prey availability.

Four groups of five female rock crabs and two groups of
five male rock crabs (32—58 mm CW) plus two groups of
five female mud crabs and four groups of five male mud
crabs (14-23 mm CW) were each presented with

approximately 200 oysters (2-9 mm SL) which were
attached to sections from chinese-hat spat collectors. The
crabs were obtained from collectors in Caraquet Bay. In
addition, six individual male and four individual female
rock crabs (50—76 mm CW) were each provided with a
section of a chinese-hat collector which held approximately
200 oysters (2—9 mm SL). After 7 days the number of
oyster spat eaten by each predator group was estimated by
counting the scars on the collector resulting from successful
acts of predation. Individual rock crabs were left 17 days
before the number of oyster spat eaten was estimated.

RESULTS

Lobsters and rock crabs appeared to encounter oysters
randomly. They would then manipulate them with their
mouthparts and anterior walking legs, and finally attempt
to crush them.

Lobsters broke small oysters (< 10 mm SL) outright
with their mouthparts or crusher chelae, whereas they
broke oysters of 10—35 mm SL with their crusher chelae
alone. The lobster's slender cutter chela grasped the oyster
while the more robust crusher compressed opposite valves
of the shell. If the shell did not break, its position was
repeatedly adjusted and further compression forces were
applied until a weak spot was found and breaking occurred.
No one part of the oyster shell appeared to be broken more
frequently, as evidenced by the varied shapes of the shell
fragments resulting from the lobster's actions (Figure 1A).
Oysters that could not be broken after several crushing
attempts were usually rejected.

Rock crabs readily crushed small oysters up to 10 mm
SL; they held the oyster with one chela and crushed it with
the other. Rock crabs appeared more specific than lobsters
in opening oysters of > 10 mm SL, although they showed a
similar propensity to "test" oysters for weak spots until
the shell ultimately broke or was rejected. The most
common approach was to chip pieces from the shell margin
until a hole was made into which the tips of the chelae
could be inserted; then, the shell valves were pried apart.
No part of the shell margin appeared to be attacked prefer-
entially. Occasionally, shell valves were not separated and a
hole or large gap was made at the edge of the shell. Oysters
of up to about 25 mm SL were also opened by making a
hole in the central area of the cup valve where the shell
was thin. Patterns of damage to oyster shells produced by
rock crabs are shown in Figure 1 B.

The two crab species exhibited different behaviors when
foraging for oysters on collectors. Rock crabs broke oysters
while they were attached or detached the oysters before
opening them. Mud crabs were restricted to breaking
attached oysters.

No observable sexual differences existed in oyster-
opening behavior for lobsters or crabs. Once the oyster had
been opened, lobsters used only their mouthparts to glean
flesh from the prey, while rock crabs used their chelae and
mouthparts to tear away the flesh from the broken shell.

Predation on Oysters by Lobsters and Crabs

131

$40^C|

o t> s) & ft a

"V ^f w Ur

50mm

Um) J

Figure 1. (A) Oyster shell fragments resulting from lobster predation; note the varied shapes of the fragments. (B) Shell fragments
from oysters opened by rock crabs; note the characteristic damage to shell margins and central areas of cup valves.

132

Elner and Lavoie

Figure 2 shows that, for all the lobster size groups, preop-
tion rates were highest within the small-to-intermediate sizes
of oysters (10—25 mm SL) and declined rapidly with larger
oysters. Rock crabs also fed on a broad size range of oysters,
but larger rock crabs preferred larger oysters than the
smaller rock crabs. Oysters of 30—35 mm SL appeared to
be at a critical size as they were rarely preyed upon by
either lobsters or rock crabs. Feeding rates were variable
among predators of the same size group and for individual
lobsters and rock crabs from day to day ;however, mean daily
predation rates increased significantly (P <0.01) as predator
sizes increased for lobsters and rock crabs (Figure 3). Thus,
maximum mean rates (± standard error of the mean. SE) by
lobsters and rock crabs, 28.0 ± 0.33 and4.5 ± 1 .40 oysters •
predator" 1 • day" 1 , respectively, were attained by some of
the larger predators of each species (Figure 3). The larger
rock crabs attained predation rates equivalent to those of
the smaller lobsters.

The mean predation rate (± SE) on attached oysters
(2—9 mm SL) by rock crabs (32—58 mm CW) in groups was
0.59 ± 0.07 oyster • crab" 1 ■ day" 1 , whereas the mean rate
for isolated rock crabs (50-76 mm CW) on attached
oysters was 1 .26 ± 0.25 oysters • crab" 1 ■ day" 1 . Mud crabs
(14—23 mm CW) in groups consumed 0.44 ± 0.09 oyster •
crab" 1 • day" 1 (Table 1). In contrast to the relationship in
Figure 3, there was no correlation (P > 0.05) between
predation rate and predator size for isolated rock crabs
feeding on attached oysters; however, the relatively larger
rock crabs held in isolation had a higher overall mean
predation rate on attached oysters than the smaller rock
crabs in groups (Table 1).

No discernable sexual differences existed in prey size
selection or predation rate for the lobsters and crabs in
any of the experimental series. The shape of the diet curves
(Figure 2) for male lobsters (58—62 mm CL) and rock
crabs (35—45 mm CW) resembled those for similar sized
female conspecifics (lobsters, 55—63 mm CL; rock crabs.
32—46 mm CW). Similarly in Figure 3, no significant
differences existed in mean predation rates between the
similar sized male and female lobsters (d. 6.93 ± 1 .96 and 9,
6.22 ± 2.57 oysters • lobster" 1 • day" 1 ; t = 0.23, df = 9,
P > 0.5) or the similar sized male and female rock crabs
(d, 0.98 ±0.11 and 9, 1 .47 ± 0.18 oysters • crab" 1 • day"';
t = 2.01, df = 10. P >0.05). Differences in mean predation
rates for male and female crabs feeding on attached oysters
were not significant (Table 1 ).

DISCUSSION

Patterns of destruction of oyster of > 10 mm SL were
specific for lobsters and rock crabs; therefore, it should be
possible to identify the predators on oyster beds by
examining oyster-shell fragments. Opening techniques
resembling those observed in our study have also been noted
by Elner and Jamieson (1979) for lobsters and rock crabs
feeding on Atlantic deep-sea scallops, Placopecten

Lobster

>>
cs
Q

C
0)

(0
UJ

n
E

C
CO

9 85 -98mm

I.Oi-

0.8

0.6

0.4

0.2

Rock Crab

9 32-46mm

cT 94 -107mm

}->£-• cf 73 -79mm \

\\ \

i 1 1 1 I

15 20 25 30 35

Oyster SL

Figure 2. Mean daily oyster consumption per predator plotted
against oyster shell length (SL) for lobster carapace length (CL) and
rock crab carapace width (CW) size groups.

Predation on Oysters by Lobsters and Crabs

133

2 30 40 50 60 70 80 90 100 110

S Carapace Length (mm)

Rock Crab

60 70 80

Carapace Width (mm)

100

110

Figure 3. Individual lobster and rock crab predation rates over the
entire range of oyster sizes eaten, expressed as mean number (± SE)
of oysters eaten per day (Y-axis) relative to predator size (X-axis).

TABLE 1.

Predation rates of rock crabs and mud crabs
on oysters (2-9 mm SL) on
chinese-hat spat collectors.

Number of

Oysters Eaten

♦over 7 days

Mean Number

Crab (mm CW)

Sex

fover 17 days

oysters

. crab . day

Rock Crabs in Groups*

34,40,50,50,58

0.49

34,38,41,43,45

0.69

32,39,46,47,51

0.34

38,40,40,41,44

0.54

37, 39,41,46,51

0.63

32,41,42,43,54

0.86

Mean daily predation rate (± SE) = 0.59

±0.10(d):0.59 ±0.11 (9)

(t =

0.02,

df=4, P>0.5)

Overall mean daily predation rate (± SE) =

0.5?

±0.07

Isolated Rock Crabsf

1.12

1.24

1.53

0.71

0.24

1.29

1.59

3.12

1.35

0.41

Mean daily predation rate (± SE) = 1.02 ±0.19 (d); 1.62 ±0.56 (9)

(t = 1.16, df = 8, P>0.1)

Overall mean daily predation rate (±SE) = 1.26 ±0.25

14, 18, 19, 20. 21
17. 18, 19, 19,23
18,18,19,20,21
16,20,23,23,23
14, 14, 14, 15, 16
16,16,17,19, 19

Mud Crabs in Groups*

d 10

d 14

d 17

d 27

9 6

9 19

(The regressions are: lobsters, Y = -15.41 + 0.37 X, R 2 = 0.48; Mean daily predation rate (± SE) - 0.49
rock crabs, Y = 0.35 + 0.02 X, R 2 = 0.29.)

0.29
0.40
0.49

0.77
0.17
0.54
: 0.10(d), 0.36 ±0.19(9)
(t= 1.28,df=4,P>0.1)
Overall mean daily predation rate (± SE) = 0.44 ± 0.09

magellanicus (Gmelin). Krantz and Chamberlin (1978)
described six distinct patterns of damage to cultchless
oyster spat by the blue crab Callinectes sapidus Rathbun;
three of the destruction patterns (crushed shells of small
oysters, chipped shell margins, and broken spat attachment
points) were the same as those observed for the rock crabs.
Our experiments showed that, notwithstanding predator
size and the proportionately smaller chelae of the rock crab,
both lobsters and rock crabs were able to feed over the same
size range of oysters. Mud crabs, also, appeared to be
effective predators for their size and were able to open
attached oysters of 2-9 mm SL at a similar rate to larger
rock crabs. Overlaps in lobster and rock crab predation on

deep-sea scallops and green sea urchins (Strongylocentrotus
droebachiensis 0. F. Miiller) have been documented pre-
viously (Elner and Jamieson 1979, Elner 1980). Thus, these
predators would compete for prey whenever they occur
together.

Considering our experimental design, where oysters of
all sizes were available, the largest oysters eaten were
probably below the absolute maximum size of oyster that
predators could open, if small oysters were unavailable.
We believe, however, that our upper limit of 30— 35 mm SL
is a realistic representation of the largest oyster size eaten
in the field where alternative prey are always present. All
lobsters and rock crabs tested were capable of opening

134

ELNER AND Lavoie

oysters of 25—30 mm SL, yet they exhibited preferences
for oysters in the 10- to 25-mm SL size range (Figure 2).
The behavior pattern of predators showing preference for
prey of less than the maximum size they can consume has
also been observed in the green crab Carcinus maeiias
(Linneaus) (Elner and Hughes 1978, Hughes and Elner 1979,
Elner and Raffaelli 1980). Size selection of prey has been
reported by Elner and Jamieson (1979) and Elner (1980)
for lobsters and rock crabs preying on deep-sea scallops and
green sea urchins. Such selection behavior can result from
the predator making an active behavioral choice based on
prey value, or a passive, mechanical consequence of prey
availability and the predator having a set "persistence time"
proportional to its hunger level (see Hughes [1980] for
review). Our observations suggest that prey size selection is
probably a passive, mechanical process. Lobsters and crabs
attempted to prey on all oysters they encountered but the
larger predators were clumsy in handling small oysters and
all predators rejected oysters if they could not break them
after a series of force applications. Thus, size-selective
mechanisms tended to shift predation pressure away from
the small (less easily handled) and large (stronger) oysters
and toward the preferred size of oysters. Both the active
behavioral and mechanical paradigms for size selection
predict that the diet curves should shift to the right as the
size of the predator increases (Hughes 1980). Although this
relationship was demonstrated for the rock crab, it was not
for the lobster.

Predictions of the impact of a predator on an oyster
stock based only on prey-selection behavior and predation
rates observed in the laboratory are not particularly mean-
ingful. Data on abundance and size frequencies of predators
and prey, as well as other factors influencing prey selection
and predation rate, are required before a realistic estimate

of predation mortality can be made. We believe, however,
that the small rock crabs and mud crabs, which are
extremely abundant on the oyster beds and spat collectors,
kill many more oysters than the more rapacious, but much
less common, lobsters and large rock crabs. Similarly,
Whetstone and Eversole (1978, 1981) have suggested that
Panopeus herbstii Milne-Edwards, a mud crab similar to
Neopanope sayi, is as important as larger crab species as a
predator of seed of the northern quahog clam Mercenaria
mercenaria (Linne) because of its relatively higher
abundance and predatory capability. Our laboratory results
support the field observations by MacKenzie (1980) that
rock crabs and mud crabs cause substantial oyster mortality
and show that the decapods tested are capable of consuming
large numbers of oysters, and thus have the potential to
reduce oyster production. Furthermore, the results show
that the 30- to 35-mm SL of oysters is a critical size at
which oysters may be virtually invulnerable to decapod
predators.

Culturists growing oysters where crabs and lobsters occur
should adjust their strategy to protect oyster seed until it
reaches 30—35 mm SL. Because small crabs can prey on
oyster spat on collectors, culturists should ensure that the
collectors are protected from invasion by these crabs.

ACKNOWLEDGMENTS

The authors gratefully acknowledge the contributions
of Messrs. M. DeGrase, R. Daigle, E. Ferguson, and the
team from the Caraquet Bay Oyster Development Plan.
Drs. V. S. Kennedy, P. Lawton, R. J. Miller, and D. J.
Scarratt gave helpful reviews of early drafts of the manu-
script. Photographs and figures were provided by Messrs F.
Cunningham and P. W. G. McMullon.

REFERENCES CITED

Elner. R. W. 1980. Predation on the sea urchin (Strongylocerttrotus
droebachiensis) by the American lobster (Homarus americanus)
and the rock crab (Cancer irroratus). Pringle, J. D., G. J. Sharp,
and J. F. Caddy, eds. Proceedings of the workshop on the rela-
tionship between sea urchin grazing and commercial plant/
animal harvesting. Can. Tech. Rep. Fish. Aquat. Sci. 954:48-65.

& R. N. Hughes. 1978. Energy maximization in the diet of

the shore crab, Carcinus maenas. J. Anim. Ecol. 47:103-116.

Elner. R. W. & G. S. Jamieson. 1979. Predation of sea scallops,
Placopecten niagellanicus, by the rock crab, Cancer irroratus,
and the American lobster, Homarus americanus. J. Fish. Res.
Board Can. 36:537-543.

Elner, R. W. & D. G. Raffaelli. 1980. Interactions between two
marine snails, Littorina rudis Maton and Littorina nigrolineata
Gray, a predator, Carcinus maenas (L.), and a parasite. Micro-
phallus similis Jagerskiold. J. Exp. Mar. Biol. Ecol. 43:151-160.

Ennis, G. P. 1973. Food, feeding and condition of lobsters, Homarus
americanus, through the seasonal cycle in Bonavista Bay, New-
foundland. J. Fish. Res. Board Can. 30:1905-1909.

Galtsoff, P. S. 1964. The American oyster Crassostrea virginica
Gmelin. U.S. Fish Wildl. Serv. Fish. Bull. 64:1-480.

Hughes, R. N. 1980. Optimal foraging theory in the marine context.
Oceanogr. Mar. Biol. Annu. Rev. 18:423-481.

& R.W. Elner. 1979. Tactics of a predator, Carcinus maenas.

and morphological responses of the prey, Nucella lapillus. J.

Anim. Ecol. 48:65-78.
Krantz, G. E. & J. V. Chamberlin. 1978. Blue crab predation on

cultchless oyster spat. Proc. Natl. Shellfish. Assoc. 68:38-41.
MacKenzie, C. L.. Jr. 1981. Biotic potential and environmental

resistance in the American oyster (Crassostrea virginica) in Long

Island Sound. Aquaculture 22:229-268.
McDermott, J. J. 1960. The predation of oysters and barnacles by

crabs of the family Xanfhidae. Proc. Pa. Acad. Sci. 34:199-211.
Menzel, R. W. & S. M. Hopkins. 1955. Crabs as predators of oysters

in Louisiana. Proc. Natl. Shellfish. Assoc. 47:177-184.
Scarratt, D. J. 1980. The food of lobsters. Pringle. J. D., G. J.

Sharp, and J. F. Caddy, eds. Proceedings of the workshop on

the relationship between sea urchin grazing and commercial

plant/animal harvesting. Can. Tech. Rep. Fish. Aquat. Sci.

954:66-91.
& R. Lowe. 1972. Biology of rock crab (Cancer irroratus)

in Northumberland Strait. /. Fish. Res. Board Can. 29:161-166.
Whetstone, J. M. & A. G. Eversole. 1978. Predation on hard clams,

Mercenaria mercenaria, by mud crabs Panopeus herbstii. Proc.

Natl. Shellfish. Assoc. 68:42-48.

. 1981. Effects of size and temperature on mud crab.

Panopeus herbstii, predation on hard clams, Mercenaria

mercenaria. Estuaries 4:153-156.

Journal of Shellfish Research, VoL 3, No. 2, 135-140, 1983.

STUDIES OF SHELL DISEASE OF THE EUROPEAN FLAT OYSTER
OSTREA EDULIS LINNE IN NOVA SCOTIA

M. F. LI 1 , R. E. DREMNAN 1 , MICHAEL DREBOT, JR. 2
AND GARY NEWKIRK 2

1 Department of Fisheries and Oceans
Fisheries and Environmental Sciences
Halifax Laboratory, Halifax

Nova Scotia, Canada B3J 2S 7

2 Department of Biology
Dalhousie University, Halifax
Nova Scotia, Canada B3H4J2

ABSTRACT Shell disease was found in the progeny of the European flat oyster Ostrea edulis Linne imported several
years ago to Nova Scotia. This disease probably accounted for fibrosis in several tissues of affected oysters, but, in general,
had no serious effect on oyster stocks in Nova Scotia. The marine fungus Ostracoblabe implexa Bornet et Flahalut was
isolated and cultured from infected shells. Electron microscopy of the organism revealed the fine structure of the ovoid
enlargements and their morphogenesis under prolonged incubation at 5 C.

KEY WORDS Oyster, Ostrea edulis, marine fungus, Ostracoblabe implexa, shell disease, oyster pathology.

INTRODUCTION

Shell disease of European flat oysters has been known
for many years, and the general symptoms of this disease
have been described in detail (Korringa 1951, Alderman and
Jones 1971b). The causative agent, however, was not
established until the isolation of Ostracoblabe implexa
by Alderman and Jones (1971a, b). The first indication of
shell disease is the development of white spots inside the
shell. As the invasion of the shell continues with the penetra-
tion of the growing mycelium, more spots appear which
coalesce to form white cloudy areas. The perforation of
the shell by the infestation appears to cause a change in the
secretions of the mantle of the host animal. The extent of
conchyolin deposition depends to great extent on the focal
intensity of the fungal attack within the shell. At the center
of the infestation the conchyolin tends to be in the form
of a wartlike excrescence which may be 2 to 4 mm in
thickness. In severe cases the excrescences become enlarged,
coalescing into one or more knobs in the muscle base.
Eventually the area of muscle attachment may become a
raised boss. This bosslike excrescence is not found outside
the muscle attachment area and typifies the disease called
maladie du pied in France.

The disease was reported in Britain, France, and the
Netherlands (Sinderman and Rosenfield 1967, Sprague 1971 ,
Alderman 1976). It was first observed in Nova Scotia in
1975 in experimental stocks of oysters transferred from
Ellerslie, Prince Edward Island. The stocks resulted from
the introduction of Dutch stocks from Milford, CT, which
were bred in quarantine. A later examination of preserved
shells showed that both the parent stock and the first
Canadian generation showed symptoms of shell infestation.
The presumptive involvement of 0. implexa was confirmed
by D. J. Alderman (Fish Disease Laboratory, Weymouth,

Dorset, England; personal communication) from examina-
tion of fresh and preserved shell material and by culture.

A study of the prevalence of the disease was carried
out by the Nova Scotia Department of Fisheries in conjunc-
tion with Dalhousie University, and the Fish Disease and
Nutrition Section of Fisheries and Environmental Sciences,
Department of Fisheries and Oceans, in the summer of
1980. The disease appeared to exist in European flat oysters
held at Whitehead Harbour and Spanish Ship Bay. This
report describes the lesions found in the infected oyster
shells, histopathological changes in the oyster tissues,
growth and isolation of O. implexa, and fine structure of
the isolated organism.

MATERIALS AND METHODS

Oysters

Hatchery-produced progeny of imported European flat
oysters O. edulis were grown on natural beds in Nova Scotia.
The 1975- to 1979-year classes of the oyster were sampled
from Whitehead Harbour or Spanish Ship Bay of Nova
Scotia in May-October of 1980. After gross examination
of the specimens for typical lesions of shell disease infec-
tion (Alderman and Jones 1971b), the shells were cleaned
thoroughly and rinsed repeatedly with sterile seawater, then
incubated in sterile seawater at 15°C. Some of the shells
were decalcified with an EDTA solution (Alderman and
Jones 1971b) or Cal-Ex® (Fisher Scientific Co., Ltd.) for
examination by phase contrast microscopy to detect the
infective agent(s) within the shell material.

Histology

For histopathological examinations tissues from 20
oysters were fixed in Davidson's fluid (Shaw and Battle

135

136

Li ET AL.

1957) embedded in paraffin, sectioned, and stained with
Harris' hemotoxylin and eosin. Photomicrographs were
prepared with a Zeiss photomicroscope.

Growth and Isolation of the Infective Agent

Fragments of shell with shell disease lesions were incu-
bated in autoclaved seawater at 15°C for 3 to 4 weeks. The
growth of fungal colonies was examined periodically. A
pure culture of the organism was obtained by incubation of
a small piece of diseased shell in a yeast/peptone medium
(Alderman and Jones 1971a, b) at 15°C. Identification of
the organism was based on morphology described by
Alderman and Jones (1971b) and Alderman (1976, 1980).

Electron Microscopy

The isolated organism was harvested from yeast/peptone
medium by low speed centrifugation. The resulting pellets
were fixed in 2% glutaraldehyde in phosphate buffer
(0.1 M, pH 7.0), postfixed in osmium tetroxide, and
embedded in TAAB® resin (Marivac Ltd., 1872 Garden
Street, Halifax, NS B3H 3R6, Canada). The ultrathin sec-
tions were stained with uranyl acetate and lead citrate
(Dawes 1971) and examined using an Hitachi HS-9 electron
microscope.

RESULTS

Figure 1 A shows white spots coalescing to form a cloudy
area on an infested shell. A lesion involving wart formation
through deposition of conchyolin by the oyster mantle is
shown in Figure IB. Figure 1C shows a heavily infested
specimen with a large sheet of conchyolin embedded in a
cloudy area of shell. Examination of several groups of

oysters indicated that the overall prevalence of this disease
was approximately 10% (Table 1).

Most of the infestations were in the early stage of
lesion development; < 1% of the infested oysters had
reached the advanced, heavy warting stage. The preva-
lence was slightly higher in the older oysters than in the
younger ones (Table 1); however, no seriously damaging
effect of this disease on the oyster stocks was observed.

TABLE 1.

Percentage occurrence of shell disease among 3- and 5-year-old

oysters during May survey (mixture of oysters from

Spanish Ship Bay and Whitehead Harbour) and

July survey (oysters from Spanish Ship Bay).

Progress of Disease (Stages)

Age

Number

3 years

1,836

132

1,990

(92.3%)t

( 6.6%)

(0.6%)

5 years

750

852

(88.0%)

(10.7%)

(0.5%)

(0.8%)

Total No.

of oysters

2,586

223

2,842

Mean % in

each class

91.0%

7.9%

0.5%

0.6%

5 years

103

Mean % in

each class

91.3%

5.8%

1.9%

1.0%

*Stage = no disease; stage 1 = one or more white spots; stage 2 =

slight warting; stage 3 = heavy warting.
fPercentages in parentheses indicate percentage occurrence of shell

disease for oysters in designated year class (Le., 3 or 5 years old).

Figure 1. Typical shell disease lesions at various stages: 1A, white spots or cloud (•<-); IB, black or brownish conchyolin at center of
the warts (•*-);!€, large sheet of conchyolin deposited in cloudy area (<-). Bar = 20 mm

Shell Disease of Ostrea edulis

137

In the tissues of 20 infested oysters examined for the
causative agent and possible histopathological changes, no
sign of an infective agent was found. Generally there was
no apparent ill effect caused by the infestation; however,
development of fibrous tissue was evident in the gill, mantle,
and digestive tracts of many of the specimens examined
(Figures 2A and 2B). Fungal mycelia were easily observed
in the decalcified specimens using phase contrast microscopy.
Formation of fungal colonies on shell fragments was usually
observed after a 3- to 4-week incubation of infested material
in sterile seawater at 15°C (Figure 3). Isolation of a pure
culture of a fungus was achieved by incubating wart tissue
in yeast/peptone medium at 15°C. Figure 4A shows the
mycelia of the isolated fungus which exhibited ovoid
swelling at frequent, irregular intervals (Alderman and
Jones 1971b; Alderman 1976, 1980). The incidence and
size of the ovoid swellings appeared to increase with incuba-
tion at low temperature (5°C) for an extended period of
time; some of the swellings appeared as spherical bodies
(Figure 4B). Figure 5 shows typical O. implexa in a decal-
cified specimen that had been incubated for 14 days at
15 C following autoclaving in seawater and inoculation
with the isolated organism.

Electron micrographs of the isolated organism are shown
in Figures 6 and 7. The mycelium contains vacuoles and
various electron-dense bodies. The organelles, such as
nucleus, mitochondria, and endoplasmic reticulum,
appeared to be well developed in the ovoid swellings
(Figure 6A-6D). When the cultures were incubated at 5°C
for a prolonged period, proliferation of the endoplasmic

reticulum was observed, and the formation of a multilayered
heavy wall often resulted (Figures 7A and 7B).

DISCUSSION

The shell lesions of infested oysters found in Nova
Scotia were typical of the shell disease described in the
literature (Alderman and Jones 1971b; Alderman 1976,
1980). The stocks sampled at Whitehead Harbour and
Spanish Ship Bay were produced in the Pleasant Point
Hatchery, where the spat and brood stocks were held in
the same tank during the first summer at an elevated tempera-
ture. The possibility that the parent stocks carried the
organism and served as a disease source cannot be ruled out.

Alderman and Jones (1971b) observed an increase in
long epithelial cells in mantle tissue of certain heavily
infested specimens. The fibrous tissue noted in infested
oysters could be a result of the shell disease infestation
because an increase in fibrous tissue formation in shellfish
appears to be a common and nonspecific reaction to
infestation or inflammation of the host animal (Pauley 1969,
Sparks et al. 1969, Sparks and Fontaine 1973). The infesta-
tion by O. implexa did not, however, appear to have any
serious physiological effect on the host animal, since most
specimens had well developed gonads and some were
spawning. Alderman (1980) suggested that levels of shell
disease infestation are high only where the water tempera-
ture exceeds 22°C for at least 2 weeks. The ambient water
temperature in Nova Scotia is generally too cold and,
therefore, precludes serious shell disease problems in oyster
stocks.

2A '___

Figure 2. Fibrous tissues of infected oysters: 2A, fibrous tissue
higher magnification (<-) (bar = 100 /im).

development in the gill (<-) (bar = 200/Llm); 2B, fibrous tissue at a

138

Li ET AL.

Figure 3. Fungal colony (<-) from an infested oyster shell; incubated
in seawater for 3 weeks at 15 C.

Figure 4. Cultures of the isolated organism in a yeast/peptone
medium at 15 C: 4 A, 7-day-old culture (note the ovoid swellings at
irregular intervals, arrowed); 4B, 4-month-old culture at 5 C (note
an increase in number and size of the enlargements of the mycelia;
some are developing into spherical chlamydosporesl^). Bar = 20 p.m

An outbreak of the foot disease occurred in populations
of the Pacific oyster Crassostrea gigas (Thurnberg) off the
Canadian west coast in the fall of 1956 (Quayle 1969).

The same organism may cause both shell and foot diseases
(Sinderman and Rosenfield 1967, Sprague 1971).
Unfortunately, the causative agent of foot disease on the
west coast was not identified.

Ostracoblabe implexa was described in relation to shell
disease of oysters almost a century ago (Bornet and Flahault
1889), but the isolation was not accomplished until 1971
by Alderman and Jones (1971a, b). One of the major
characteristics of the organism is the presence of ovoid or
spherical swellings in the mycelium (Alderman and Jones
1971b; Alderman 1976, 1980). Our isolate showed similar
ovoid enlargements and fine structure of the prochlamydo-
spore as described in the literature. Our results further
demonstrated the morphogenesis of the chlamydospore
during incubation at low temperature. No sexual reproduc-
tive phase was observed. Ostracoblabe implexa has been
placed in the phycomycetes but its exact taxonomic
position remains to be determined.

Figure 5. An experimentally infested oyster shell (SH) incubated in
seawater for 3 weeks at 15°C. Phase contrast of specimen decalcified
by Cal-Ex®. Note the typical ovoid swellings (•«-). Bar = 20 flm

ACKNOWLEDGMENTS

We thank the Nova Scotia Department of Fisheries for
funding part of this study at Dalhousie University; Dr. D. J.
Alderman for his examination of shell samples; Robert
Zwicker and John Cornick of the Fish Health Unit, Disease
and Nutrition Section, for supplying some of the oyster
specimens; Dr. D. Brewer at the Atlantic Research Labora-
tory of the National Research Council of Canada for his
constructive discussion of the taxonomic position of the
isolated organism; Dr. L. E. Haley of Dalhousie University
and Mr. T. Rowell of Invertebrates and Marine Plants
Division for reviewing the manuscript and for their criticisms;
and Ms. Vivian Marryatt for her technical assistance with
the histological examinations.

Shell Disease of Ostrea edulis

139

IW
OW r

er- ^r

6B ^

JDW

aOW

^iw

Figure 6. Electron micrographs of mycelium and prochlamydospores from an isolate grown in a yeast/peptone medium for 7 days at 15 C:
6A, mycelium containing vacuoles and electron-dense bodies; 6B and 6C, ultrathin sections of prochlamydospores; 6D, cross section of
prochlamydospores. Note the spores containing inner and outer walls, nucleus, vacuoles, mitochondria, and numerous electron-dense bodies.
(N, nucleus; OW, outer wall; IW, inner wall; V, vacuole; ED, electron -dense bodies; M, mitochondria; ER, endoplasmic reticulum) Bar = l^lm

140

Li ET AL.

-MW

n - »■*

Figure 7A and 7B. Chlamydospores from a culture which had been incubated at 5 C for 4 weeks. Note the curling arrangements of micro-
tubules and development of a thick, multilayered wall. (N, nucleus; V, vacuole ;MW, multilayered wall; ER, endoplasmic reticulum) Bar = 1 [lm

REFERENCES CITED

Alderman, D. J. 1976. Fungal diseases of marine animals. Jones,
E. B. G., ed. Recent Advances in Aquatic Mycology. London,
G.B.: Paul Elek (Scientific Book) Ltd. 749 p.

. 1980. Shell disease of oysters. Diagnostic summaries of

diseases of fish, Crustacea and molluscs by working group on
pathology of marine animals. Int. Counc. Explor. Sea 91-94:00.

& E. B. G. Jones. 1971a. Physiological requirements of two

marine Phycomycetes, Althornia crouchil and Ostracoblabe
implexa. Trans. Br. Mycol. Soc. 57(2):213-225.
. 1971b. Shell disease of oyster. Fish. Invest. Ser. II Mar.

Fish. G.B. Minst. Agric. Fish. Food 16(8) :20 p.
Bornet, E. & C. Flahault. 1889. Sur quelques plantes vivant dans le

test calcaire des mollusques. Bull. Soc. Bot. Fr. Ser. 2:11 p.
Dawes, C. J. 1971. Biological Techniques in Electron Microscopy.

New York, NY: Harper and Row Publishers, Inc. 193 p.
Korringa. P. 1951. Investigation on shell-disease in the oyster,

Ostrea edulis L. Rapp. P. B. Reun. Cons. Int. Explor. Mer.

128(2):50-54.

Pauley, G. B. 1969. A critical review of neoplasia and tumor-like

lesions in mollusks. Natl. Cancer Inst. Monogr. 31 :509— 539.
Quayle, D. B. 1969. Pacific oyster culture in British Columbia.

Bull. Fish. Res. Board Can. 169:1-193.
Shaw, B. L. & H. I. Battle. 1957. The gross and microscopic anatomy

of the digestive tract of the oyster Crassostrea virginica (Gmelin).

Can.J.Zool. 35:327-347.
Sindermann, C. J. & A. Rosenfield. 1967. Principle diseases of

commercially important marine bivalve Mollusca and Crustacea.

U.S. Fish Wildl. Serv. Fish. Bull. 66:335-385.
Sparks, A. K. & C. T. Fontaine. 1973. Host responses in white

shrimp, Penaeus setiferus. to infection by the larval trypanor-

hynchid cestode, Prochristianella penaei. J. Invertebr. Pathol.

22:213-219.
Sparks, A. K., G. B. Pauley & K. K. Chew. 1969. A second

mesenchymal tumor from a Pacific oyster (Crassostrea gigas).

Proc. Natl. Shellfish. Assoc. 59:35-39.
Sprague,V.1971. Disease of oysters./lnn. Rev. Microbiol. 25 : 21 1-230.

Journal of Shellfish Research, Vol. 3, No. 2, 141-151. 1983.

THE ORIGIN AND EXTENT OF OYSTER REEFS IN
THE JAMES RIVER, VIRGINIA 1

DEXTER S. HAVEN AND JAMES P. WHITCOMB

Virginia Institute of Marine Science
and School of Marine Science
The College of William and Mary
Gloucester Point, Virginia 23062

ABSTRACT The public oyster grounds (Baylor Survey Grounds) in the James River, VA. were studied with respect to
bottom type and oyster density from 1978 to 1981. Approximately 10,118 ha (25,000 acres) were investigated using an
electronic positioning system to establish station locations. Bottom types were determined using probing pipes, patent
tongs, and an acoustical device. About 17.1% of the bottom was classified as consolidated oyster reef, and 47.5% was
moderately productive mud-shell or sand-shell bottoms. The remaining 35.4% was rated as unsuitable for oyster culture.
The surface configuration of oyster reef areas in the James River is similar to those in coastal lagoons along the Gulf of
Mexico. They are thought to have developed in the James River as they did in the Gulf of Mexico area as sea level rose
during the Holocene Period.

KEY WORDS

INTRODUCTION

The naturally productive oyster-growing areas in Virginia
were surveyed and set aside for public use in 1 894 by Lt. J. B.
Baylor (Baylor 1894) and since then have been designated
as Baylor Grounds. Statewide, they comprise about 98,324 ha
(243,000 acres) with 10,118 ha (25,000 acres) located in
the James River, VA (Haven et al. 1981a). The Baylor
Survey outlined only broad areas of naturally productive
bottoms and did not delineate nor quantify the size or
shape of individual oyster reefs. Consequently, many unpro-
ductive areas (mud and sand bottoms) were included within
the bounds of the survey (Moore 1911, Loosanoff 1931,
Haven et al. 1981a).

This paper describes and quantifies the seed-oyster
producing regions in James River, VA, within the bounds of
the public (Baylor Survey) oyster grounds. It is a portion of
a much larger investigation which evaluated the suitability
for oyster culture of nearly all public oyster grounds in
Virginia (Haven et al. 1981b). The area studied, divided
into five zones, is shown in Figures 1 and 2.

Prior to this study there were only two attempts to
quantify productive and nonproductive areas within the
Baylor Grounds. The first was conducted in 1910 using a
chain drag, hand tongs, and a lead line to outline bottom
types and quantify oyster density (Moore 1911). Positions
were established by sextant bearings and about 10,440
soundings were taken. A second study was conducted
between 1973 and 1976 which demonstrated significant
changes in oyster density along seven corridors in the James
River, but the area of the various bottom types were not
determined (Loesch et al. 1975).

Contribution No. 1199 from the Virginia Institute of Marine Sci-
ence, The College of William and Mary, Gloucester Point, VA 23062.

The James River has been and continues to be of major
importance to the oyster industry in Virginia. Oysters set
and survive well there but growth is slow and meat quality
is typically poor (Loosanoff 1931, Haven et al. 1981b).
Since the mid-1 800's, small oysters of less than 7.6 cm
(3 in.) in length (termed seed oysters) have been harvested
from the river and transplanted to other areas where
growth and meat quality improved. In the past 50 years,
an estimated 75% or more of the seed oysters planted in
Virginia by private interests on leased bottoms came from
the James River (Haven et al. 1981b).

From about 1920 to 1945 annual seed-oyster production
in the James River averaged about 1,675,000 Virginia
bushels (82,346 m 3 ) (Marshall 1954), and from 1946 to
1961 it averaged between 1.5 to 2.5 million (73,800 to
123,000 m 3 ). Between 1961 and 1981, however, yearly
production fell drastically and in that period it fluctuated
between 250.000 and 550,000 bushels (12,300 and
27,075 m 3 ) (Haven et al. 1981b).

The decline in landings has been associated in part with
a decline in demand for seed oysters because of the impact
of the oyster pathogen Haplosporidium nelsoni (Haskin,
Stauber and Makin), commonly called MSX, on adult popu-
lations growing in high salinity waters (Haskin et al. 1966.
Andrews 1968). An additional cause of the decline in seed
production was the low demand for seed resulting from
unfavorable economic conditions such as high growing
costs and an unstable market for the final product (Haven
et al. 1981b). Accompanying the decline in landings was a
decline in spatfall intensity which was most severe in the
lower half of the seed area (Haven et al. 1981b, Andrews
1982) (Table 1). The cause of this latter decline has not yet
been adequately explained. The James River, like most of
Chesapeake Bay, has in the past three decades experienced

141

142

Haven and Whitcomb

LAWNES PT

JAMES RIVER

BLUNT PT.
*\ V

Figure 1. Oyster reefs and other bottom types in the James River, VA. Shown are areas I, II, and III separated by the clear lines and transects
A, B, C, and D. Mud bottoms within the bounds of the Baylor areas are unstippled.

Origin and Extent of Oyster Reefs in James River

143

Meters

Nautical Mile

f.:: : .;-::"j Shell and Mud
Sondand Shell

IOOO

Boylor Line

Figure 2. Oyster reefs and other bottom types in the James River, VA. Shown are areas IV and V separated by the
clear lines. Mud bottoms within the bounds of the Baylor areas are unstippled.

144

Haven and Whitcomb

increased levels of nutrient enrichment, toxic chemicals,
sedimentation, and other human alterations (Haven et al.
1981b), all of which may have affected setting of spat.

TABLE 1.

Mean spatfall per Virginia bushel of bottom substrate
at representative locations from 1947 to 1980.*

Point

Deep Water

Period

Brown Shoals

Wreck Shoals

of Shoals

Shoals

1947-1950

718

1901

385

1744

1951-1955

1030

1945

336

872

1956-1960

412

995

—

468

1961-1965

298

135

113

1966-1970

249

334

1971-1975

167

1976-1980

199

169

534

*1947-1965 data from Andrews (1982).

Hydrography of the James River

The hydrography of the James River has been the subject
of several major studies but many details are still poorly
understood. Basically, it is a partially mixed tidal estuary
(Pritchard 1953, Nichols 1972b); recent studies suggest it
may undergo a cyclic stratification-destratification process
related to the neap and spring tidal cycles (Haas 1977).

Published information on salinity from 1949 to 1961 at
Deep Water Shoals showed a range from about 2 to 10 ppt,
at Wreck Shoals from 7 to 14.5 ppt, at Newport News Point
from 12.5 to 18.5 ppt, and at Nansemond Ridge from 13.5
to 19.5 ppt (Table 2). Additional data for all stations
from 1963 to 1981 showed a similar range (VIMS unpub-
lished). Freshets occur at irregular intervals in this estuary
and 0.0 ppt has been recorded as far downriver as Wreck
Shoals (Andrews et al. 1959, Haven et al. 1976). Salinities
of 0.0 ppt commonly occur at Deep Water Shoals where
oysters are frequently killed by fresh water in the spring
of the year (Andrews et al. 1959).

TABLE 2.

Mean salinities (in ppt) in the James River, VA,
from 1949 to 1961.*

Stations

Season

Deep Water

Shoals Wreck Shoals

Newport News
Point

Nansemond
Ridge

Spring

2.0

7.0

12.5

13.5

Summer

10.0

14.0

17.5

18.5

Fall

5.0

14.5

18.5

19.5

Winter

—

13.0

16.0

16.5

*Adapted from Stroup and Lynn (1963).

The natural channel in the lower James River lies close
to the north shore, near Newport News Point, and toward
the south shore in the Burwell Bay area. In the upper

estuary near Deep Water Shoals, it is near the center of the
river. Rocklanding Shoals Channel was cut through the
northern edge of the seed areas and its depth in 1976 was
7.6 m (25 ft) (Figure 1 ).

The names of individual seed areas in the James River
have remained virtually unchanged for over 100 years.
For example, the oyster reef known as Deep Water Shoal,
marks the upriver limit of commercial production and
Nansemond Ridge is the lower limit (Figures 1 and 2).
These names can only be used to designate the general
location of a seed-producing area because one area grades
imperceptibly into another.

MATERIALS AND METHODS

The criterion for defining the naturally productive areas
is based on one aspect that is considered of major impor-
tance. The naturally productive areas in the James River
(those having oysters or shells) have existed in nearly the
same location since 1854 (Moore 1911. Marshall 1954).
Moreover, as will be discussed later, many probably existed
in the same approximate location for much longer periods
as was determined for Gulf of Mexico oyster beds (Bouma
1976). This study was designed to detect shells or living
oysters in or on the bottom. Their presence was indicative
of productive or previously productive bottoms.

The survey vessel was navigated at a speed of about
5.5 km-h -1 (3 knots) within the bounds of Baylor Grounds
along a series of transects which were delineated using the
Raydist® (manufactured by Teledyne Hastings Corp.,
Hampton, VA) electronic positioning grid system with a
precision of ± 2 m. While traversing these transects, the
bottom was probed with a 2.5-cm diameter copper pipe
every 60 to 90 m to determine bottom type. The probing
interval was decreased when the bottom type changed
rapidly. Transects were usually about 183 m apart. Studies
on bottom types were completed during 1979; sampling
for oyster density was carried out in 1981.

The presence or absence of shells and/ or oysters between
probe stations was monitored continuously with an under-
water microphone mounted in a steel frame and dragged on
a cable about 37 m behind the vessel. The sounds made by
the microphone bouncing over shells or oysters or sliding
over sand or mud were amplified and broadcasted. The
intensity and frequency of the sounds and the percentage
of time the microphone was impacting on shells or oysters
or other bottom types between stations were recorded by
the operator (Haven et al. 1979). Depths were monitored
continuously with a recording fathometer. These latter
readings were used to reconstruct four longitudinal profiles
across various bottom types.

For each station, Raydist® coordinates, coded informa-
tion on bottom types obtained with the probe, acoustic
information, and depths were recorded on tape using a
Teledyne/Hastings printer. Later, the data on the printed
tape were plotted on a series of 1:10,000 charts The

Origin and Extent of Oyster Reefs in James River

145

charts showed latitude and longitude. 1.8- and 5.5-m (6-
and 18-ft) depth contours, outlines of the shorelines, out-
lines of the Baylor Grounds, and information on bottom
types. Subsequently, the boundaries of the various bottom
types were outlined on the charts. Areas of various bottom
types were determined with a digitizing planimeter.
The following bottom types were described:

Oyster reef: firm bottom, probe penetrated to 5 cm. Shells
and oysters were typically abundant. Shells or oysters were
detected using the microphone from 75 to 1007© of the time
between the probe stations.

Sand-shell: The firm bottom consisted largely of unconsoli-
dated shell; probe operator detected the gritty texture of
sand. Shells or oysters were detected using the microphone
from 25 to 75% of the time.

Mud-shell: The probe operator detected a moderately firm
crust over a soft bottom. The probe, after penetrating the
crust, could be thrust at least 0.2 to 0.6 m further into the
bottom. Unconsolidated shells or live oysters were usually
detected using the microphone from 25 to 75% of the time
between stations.

Mud: On these soft bottoms the probe could often be
pushed almost 1 m into the bottom with little effort. They
consisted largely of mixtures of silts and clays with some
sand (Nichols 1972a). Shells and oysters were usually absent,
or very few as determined using the microphone.
Sand: These were firm bottoms, and the probe typically did
not penetrate more than 2 cm. Few shells or oysters were
detected using the probe or underwater microphone. Probe
operator detected gritty texture of sand.

After the bottom types were outlined on charts, the
bottoms in Areas II and III (Figure 1) were sampled with
hydraulically operated patent tongs. Each tonggrab sampled
an area of 0.68 m 2 (7.29 ft 2 ) and penetrated the bottom
about 10 cm on oyster reef and 30.5 cm on mud bottoms;
each sample consisted of at least one-half of a Virginia
bushel (one Virginia bushel = 0.05 m 3 ). A total of 476
sampling stations were randomly chosen along transects
defined using the Raydist® system. Data from each grab
were recorded as follows: numbers and volumes (in U.S.
quarts where 1 quart = 0.91 liter) of oysters exclusive
the current year's spat, volume in quarts of shells and
fragments, and estimates of the percentage of unburied
shell as identified by the presence of fouling organisms.
These data were used to calculate oyster density (number •
m ) and the percentage of each grab that was composed
of shells and shell fragments.

A preliminary analysis of data on oyster density indi-
cated a skewed distribution with a high percentage of zero
values; therefore, densities were analyzed for possible
significant differences in modal values using the Mann-
Whitney test for nonparametric data (Sokal and Rohlf 1981).
Oyster distribution obtained in this study was compared to
distribution found in 1910 by Moore (1911 ).

National Oceanic and Atmospheric Administration
(NOAA) charts 12248 and 12222 (1:40.000) were used in
this study to outline depth contours and shorelines. Because
these charts show depths in feet and distances in nautical
miles, these same units are used to delineate depth contours
and distances shown in the illustrations and in some of the
tabular material. In the text the following conversions are
used: the standard 6- and 18-ft contour depths are 1.8 and
5.5 m, respectively. One nautical mile (6,000 ft) is equal
to 1.83 km.

RESULTS

Reef Areas

Areas classified as oyster reef show distinctive outlines in
different parts of the estuary. In Area I six small reefs
existing near the channel are generally elongate and parallel
to the axis of the estuary and to the currents. They occur
at depths ranging from 1.8 m to more than 5.5 m (Figure 1).

Area II is characterized by larger oyster reefs, most of
which differ in shape from those in Area I (Figure 1). On
the northeastern side of Rocklanding Channel, they begin
about 1 .4 km offshore (beyond the 1 .8-m contour) and
extend to Rocklanding Channel. Many are extensive and
appear to be oriented parallel to the current and the axis of
the river. Usually, however, there is an almost equal
component oriented at right angles to the shore and the
current. A similar type of orientation exists on the exten-
sive reef area along the southwestern side of Rocklanding
Channel. There the reefs extend to the south for a maxi-
mum distance of about 3.7 km, at depths ranging from 1.8
to 5.5 m (Figure 1 ).

The oyster reefs in Area III are among the most produc-
tive in James River, and Rocklanding Shoal Channel passes
through the center of this area. On the northeastern side of
the natural channel (off Lands End) between the 1.8- and
5.5-m contour intervals, the oyster reef areas form well
defined and approximately parallel rows which are approxi-
mately at right angles to the axis of the river (and current).
Frequently, a reef ends as an isolated series of small reefs
still in line with the larger one. On the southwestern side of
the estuary in Area III, the oyster reefs are irregular in
outline but the trend appears to be parallel to the channel
as in Area I. Many are located at depths of less than 1 .8 m.
This is in contrast to the distribution noted on the north-
eastern side where most occur between the 1.8- to 5.5-m
contour lines (Figure 1 ).

In Area IV on the northeastern side of the natural
channel, which varies in depth from about 7.3 to 15.8 m,
irregularly shaped reefs occur between the 1.8- and 5.5-m
contours (Figure 2). Here, in contrast to the upriver areas,
there is no apparent orientation with respect to the axis of
the river (Figure 2). On the southwestern side, the depths
of the reef areas differ from those on the opposite side
because they exist primarily in less than 1.8 m of water.

146

Haven and Whitcomb

They are, however, similar in that they have no apparent
orientation.

Oyster reefs in Area V (Figure 2) are usually small and
scattered and are oriented at right angles to the axis of the
river and are, therefore, similar in this respect to those in
Areas I and II. Moreover, they are usually at depths less
than 1.8 m as are most reefs on the southwestern side of
this estuary.

Other Bottom Types

In Areas I through IV, sand-shell bottoms generally occur
inshore of oyster reef areas and often extend into the
inshore margin of Baylor Grounds; in Area V, where sand-
shell bottoms are scarce, they occur largely between the
reefs. Areas of mud-shell are the most extensive bottom
type in Areas II, III and IV and they occur offshore of
sand-shell bottoms. Oyster reefs in all zones are usually
surrounded by this type of bottom.

Sand bottoms are not common in the James River
Baylor Grounds; when they do occur, they are generally
located inshore of sand-shell areas. Mud bottoms are
extensive and occur in all five segments as large irregular
zones between shelled areas and in the deeper channels
(Figures 1 and 2).

Acreage of Subaqueous Bottom Types

Mud-shell bottoms were the most extensive and totaled
29.8% (3,030 ha) of the Baylor Grounds surveyed
(10,178 ha). Oyster reefs and sand-shell are about equally
abundant and comprise 17.1% and 17.7% (1, 744 and
1,800 ha), respectively, of the total area. Therefore, about
64.6% (or 6,574 ha) of the Baylor Grounds in the James
River can be classified as productive or potentially produc-
tive (Table 3).

The nonproductive mud, sand, and buried-shell bottoms
make up 35.4% (3,604 ha) of the total 10,178-ha area.
These latter types have little, if any, potential for oyster
culture.

Oyster and Shell Densities

Patent-tong sampling showed a wide variation in oyster

density on the various types of bottom. This was expected
because a previous study during 1973 and 1974 showed
that oyster distribution in the James River was typically
noncontiguous (Loesch et al. 1975). The present study
showed that oyster densities on all bottom types ranged
from to 274 oysters*rrf 2 (Table 4). Oyster-reef bottoms
had the highest mean density and ranged from a mean of
34.8'irf 2 in Area II to 28.0'irf 2 in Area III. Sand-shell and
mud-shell bottoms supported about 50 to 75% fewer
oysters. No oysters were recovered in eight samples taken in
Area II on mud and sand bottoms. On similar substrates in
Area III, oyster densities ranged from 2.2 to 10.7-irf 2 . This
latter value, discussed later, seems atypical.

A statistical analysis using the Mann-Whitney test for
nonparametric data (Sokal and Rohlf 1981) showed that
the modal grouping for oyster density (Table 4) on oyster-
reef areas was significantly higher than for mud-shell and
sand-shell bottoms in Area II (Table 5). Mud-shell bottoms
have a significantly higher modal grouping than sand-shell.
No oysters were found on sand or mud bottoms (Table 4).

In Area III, oyster-reef bottoms have a modal grouping
of oyster densities higher than all bottom types tested
(Table 5). Sand-shell bottoms were significantly higher
than mud-shell, and both have a modal grouping higher
than sand. Mud bottoms seemed to show anomalous situa-
tions because oyster densities were higher than those found
for sand-shell bottoms. A possible reason for this will be
covered in the Discussion section.

Analysis of the patent-tong data showed that bottoms
classified as oyster reef (on the basis of data obtained using
a probe and sonic gear) also contained the highest content
of shell material. In Areas II and III, shells and fragments
averaged from 42.8 to 33.9%, by volume, respectively, of
the grab's content. The high shell content and high values
for oyster density are responsible for the firmness of
bottoms classified as oyster reef. In addition, almost half
of the shell material on oyster reef bottoms was surface
shell which was exposed to the flow of the current
(Table 6).

Bottoms that were classified as mud-shell or sand-shell
in Areas II and III differed from oyster reef bottoms

TABLE 3.
Areas of various types of bottom in the James River, VA, expressed as hectares and as percent of total in each of the subareas (I-V).

Total Area (ha)

Size of Each Bottom Type (% Total) in Each Subarea

Percent Total

Bottom Type

ItoV

III

All Areas

Oyster Reef

1,744

5.1

28.0

14.1

28.5

2.8

17.1 )

Sand-Shell

1,800

35.8

22.6

16.5

5.5

19.9

17.7

64.6

Mud-Shell

3,030

14.5

29.7

33.5

31.3

23.7

29.8 j

Sand

623

11.6

4.6

6.2

1.5

10.5

6.1 |

Soft Mud

2,811

33.0

15.1

29.7

32.8

34.8

27.6

35.4

Buried Shell

170

< 0.1

0.4

8.3

1.7 j

Total hectares

10,178

298

2533

3903

1466

1978

Origin and Extent of Oyster Reefs in James River

147

because they had smaller volumes of shell material and
lower percentages of surface shell; they were less consoli-
dated and more scattered.

TABLE 4.

Density of oysters collected with patent tongs in
the James Rivei seed area.*

Area II

Area III

Bottom Types

N Mean

Range

Mean

Range

Oyster Reef

19 34.82

to 165.76

27.98

to 273.81

Sand-Shell

27 9.0

to 109.52

6.48

to 35.52

Mud-Shell

19 13.40

Oto 118.90

188

5.75

to 59.20

Sand

2.18

to 41.44

Mud

10.72

to 112.48

*From Statistical Summary of Means and Range ( 1 98 1 ).

TABLE 5.

A statistical comparison using the Mann-Whitney test of modal

grouping of oyster density (m ) in Areas II and HI in the

James River, VA. (Mean values for numbers of oysters

per m are shown in Table 3.)

Bottom Type

Levels of Significance

Oyster reef versus

mud-shell
Oyster reef versus

sand-shell
Mud-shell versus

sand-shell

Oyster reef versus

mud-shell
Oyster reef versus

sand-shell
Oyster reef versus

sand
Mud-shell versus

sand-shell
Mud-shell versus

sand
Sand-shell versus

mud
Mud versus sand
Mud-shell versus mud

Area II

Difference significant at 0.25 >P >0.01

Difference significant at 0.01 >P >0.001
Difference significant at P = 0.01

Area III
Difference significant at P < 0.001

Difference signfiicant at P < 0.001

Difference significant at P <C 0.001

Difference significant at 0.01 >P >0.001

Difference significant at 0.05 >P >0.02

Difference significant at 0.01 >P > 0.001

Not significant at P = 0.10
Not significant at P = 0.10

Transects

Elevations and slopes were studied across the oyster
reefs, or shoals, on four transects in the area near Point of
Shoals Light (Figures 1 and 3). Those transects crossed
productive oyster reefs such as Wreck Shoal and Point of
Shoals. The overall slope from the channel to the sandy
margins along the shore ranges from about 0.04 to 0.1 1 m
(0.13 to 0.35 ft) vertically for each 30.5 m (100 ft)

horizontal distance (slopes: 1:769 to 1:286, respectively).
Frequently, the elevation of the bottom from a nonproduc-
tive slough to a productive shelled area was less than 0.30 m
(1 ft) vertically for every 30.5 m (100 ft) horizontally.
Very steep slopes occur adjacent to the channel or mud
sloughs where they join productive oyster-reef or mud-shell
substrates. These sharp slopes may be as large as4.6m(15 ft)
vertically in 30.5 m (100 ft) horizontally (a slope of 1 :6.7).
Sand-shell bottoms occur as flat areas and are usually near
the shore.

DISCUSSION

Samples obtained with patent tongs in Areas II and III
confirmed observations made using a bottom probe, acoustic
gear, and fathometer. Oyster reef bottoms had higher
densities of oysters and shell material. Sand-shell and
mud-shell bottoms had lower densities of oysters and shells.
Sand bottoms seldom contained shells or oysters. Mud
bottoms, while definitely soft, sometimes contained signifi-
cant numbers of oysters.

The surface outlines of oyster reefs in the James River
may be separated into four types which closely resemble
those that occur in lagoonal systems of the Gulf of Mexico
(Graves 1905, Hedgpeth 1953, Price 1954, Scott 1968,
Bouma 1976). The longitudinal type, for example, is repre-
sented in the James River by those shown on Area I where
tidal currents are rapid over shoal bottoms. The large
irregular type is common throughout the estuary and has
two components; one is at a right angle to the axis of the
river and a second is parallel to the axis (Area II). A third
type, termed a transverse reef, is long and lies at right
angles to the current as seen in Area II off Lands End
(Figure 1). The last type, without any obvious shape, is
termed a pancake reef (Scott 1968); these are common in
Area V (Figure 2).

While those bottoms that were classified as sand-shell
and mud-shell in the James River support live oysters and
are moderately productive, we do not believe them to be
long-term features of the estuary at specific locations as
are oyster reef areas. This concept was originally discussed
by Moore (1911) who stated that the boundaries of the
highly productive areas in the James River seed area, which
approximate our oyster reef classification, were originally
sharply marked and separated from the barren (mud or
sand) bottoms. Moore (1911) speculated that operations by
man (harvesting activities and culling of the catch) over the
years were responsible for scattering shells and oysters
between the reefs and onto otherwise barren bottoms. The
atypical value of 10.7 oysters*m~ 2 on mud bottoms shown
for Area III (Table 4) probably resulted from this activity.

Oysters do not grow or survive well on sand and mud
bottoms because of several physical factors. Mud bottoms
in the James River are areas of active sedimentation (Nichols
1972a); in that environment, oysters may be covered with
sediment faster than they can grow (MacKenzie 1983).

148

Haven and Whitcomb

TABLE 6.

Number of oysters per m , exclusive of 1979 spat set, and amounts of surface and buried shells on
five bottom types in the James River, VA (August 1979).

Bottom Type Number Sampled Mean Number • m

Percent Shell

Percent Surface Shell

Percent of Sample
with Surface Shell

Area II

Oyster reef

Sand-shell

Mud-shell

Sand

Mud

Oyster reef

Sand-shell

Mud-shell

188

Sand

Mud

34.8

42.8

9.0

23.1

13.4

16.0

0.0

12.0

0.0

5.1

Area III

27.98

33.9

6.48

23.1

5.75

11.8

2.18

9.9

10.72

6.8

47.7

16.1

17.9

0.0

41.8

25.0

13.2

8.1

8.5

94.7

48.1

36.8

0.0

90.1

81.0

41.0

9.0

8.0

■

ROCK

■

MUD-SHELL

SAND-SHELL

□

SAND

□

MUD

CHANNEL

10
20
30-

DAYS PT SHOAL

WRECK SHOAL

1 1 ' '

10,000
DISTANCE IN FEET

-. 1 1 1 " 1 1

15,000 20,000

~i 1 1 1 1 1 1 r

5000

Figure 3. Longitudinal profile of various oyster bottom types along transects A, B, C and D (see Figures 1 and 2).

Origin and Extent of Oyster Reefs in James River

149

Sand bottoms, while firm, offer an unstable, shifting
substrate and sand grains are abrasive and difficult to void
from the mantle cavity when washed in by wave or current
forces. We speculate that conditions for recruitment and
growth on mud-shell or sand-shell areas may often be
marginal or they may fluctuate to a greater degree than
oyster reef areas.

The extent and depth of buried oyster shell deposits
below the reefs in the James River are not known; however,
about 2.0 X 10 6 m 3 of buried oyster shells were dredged
commercially between 1963 and 1969 from the southern
side of this estuary approximately 6 km southwest of
Newport News Point (Figure 2) (Va. Comm. Fish. Rept.
1969, Haven et al. 1981b). An early study of lagoonal
systems in the Gulf of Mexico showed that exposed oyster
reefs often extended down into the sediments for at least
2.7 m (Norris 1953). Later Bouma (1976), working in the
same area, related reef oyster formation to the world-wide
rise in sea level during the Holocene Period (Emery and
Uchupi 1972). He concluded that most of the present-day
oyster reefs in San Antonio Bay exist on top of old reefs
that started to grow about 9.000 years ago in the former
river cuts incised in late Pleistocene deposits as the sea level
began to rise. He demonstrated that shell deposits extended
as deep as 21 m (69 ft) below the sediment surface and his
14(- data showed ages of buried shell from 1,500 to 9,000
years. Bouma (1976) also stated that many surface reefs were
probably connected or adjacent to buried shell deposits.

The James River Basin and Gulf of Mexico areas
experienced the same rise in sea level during the Holocene
Period. In relation to this event, the James River Basin
flooded with seawater between 9,000 and 6,500 years ago.
The original flooding occurred along the axis of the river as
defined by the deeper channels that today range in depth
from 8 to 29 m (Nichols 1972a). The sea level has increased
about 0.6 m in the James River between 1854 and 1954.

It has yet to be determined how far oyster reefs extend
into bottom sediments in the study area; however, on the
basis of similarity in shape of oyster reefs in the James
River and Gulf of Mexico areas and the similar geological
histories, we speculate that oyster reefs in the river are
underlain with shell deposits of varying thickness and that
the reefs evolved as they did in the Gulf areas from old
shore or bottom features as sea level rose.

There have been slow changes in water depth over
oyster reefs in the James River over the last century.
Marshall (1954), using depth data from U.S. Hydrographic
charts from 1854-55 to 1943-48, stated that considerable
variations existed in the physiographic changes in the
surfaces of the seed beds (tops of the oyster reefs) during
that period. At most points depth comparisons over the
100-year period, after allowing for the increase in sea level,
indicated a decline in elevation of about 0.18 m (0.6 ft).
He speculated that this decline was the net effect of both
natural phenomena and fishery activities.

Our data, when compared with those obtained by Moore
in 1910 (Moore 1911), suggest no major differences in
oyster density in 1911 and 1981. Moore reported oyster
densities for about 590 locations in the seed area and used
them to separate bottoms into five classes (Table 7). Those
classifications were a combination of numerical data on
oyster density coupled with Moore's concept of how many
oysters a waterman needed to harvest during a 9-hr day at
the former price of $0.20 to $0.30/bu for seed and $0.45/bu
for market oysters. Certain of his categories are still valid.
Moore's barren category is comparable to our mud or sand
classifications; both have a very low potential for growing
oysters. Moore's dense growth is equivalent to our oyster
reef classification, and our definition of productive bottoms
(oyster reefs and mud-shell or sand-shell bottoms) is com-
parable to Moore's dense, scattered, very scattered and
depleted categories (Table 7).

TABLE 7.
Classification of oyster bottoms in the James River, VA.*

Oyster Harvest in Virginia Bushels
by a Tonger in a 9-hour Day

Oyster Density

Seed Oysters

Market Oysters

Barren (no shell or oysters)

Depleted

Very scattering (scattered)

Scattering (scattered)

Dense

4
4- 8
8-12
12

3
3-5
5-8

"Classification from Moore (1911).

Using the preceding categories, the following comparisons
are made (Table 8). In 1910 (Moore 1911), mean oyster
densities on dense bottoms ranged from 26.9 to 35.4 oysters-
m" 2 in Area II. In contrast, our randomly collected reef
samples in 1981 showeda similar density of 34.8-nT 2 . Mean
oyster densities on scattered to depleted bottoms in Moore's
study (1911) ranged from nearly zero to a maximum of
20.2 -m~ 2 while mean densities for comparable bottom
types in 1981 ranged from 9.0 to 13.4*m" 2 . In Area III,
three stations in Moore's study ranged in density from
32.9 to 57.0-m~ 2 ; our mean density for oyster reefs in
the same general area was 28.0ttT 2 . Mean densities in areas
of scattered to depleted bottoms ranged from zero to
33.1 -nf 2 in the early 1900's; our density data showed a
mean range of 2.2 to 10.7-irf 2 (Table 7). The overall
similarities in density for dense and reef bottom types
were unexpected because of the decline in setting intensity
in the James River that began in 1960 (Haven et al. 1981b).
We speculate that, in 1910. the intense harvest may have
depleted the beds to low levels, even when oysters were
setting at a much higher rate.

150

Haven and Whitcomb

TABLE 8.
Mean densities of oysters on various bottom types in the James River, VA, 1910- 1981. (Locations shown in Figure 1.)

1910 (Moore 1911)

1981 (Present Study)

Oyster Reefs

Growth Type

Oysters/m

Location

Substrate

Oysters/m 2

Area 11

Horse Head

Dense
Scattering
Very Scattering
Depleted

35.4

15.4

20.2

0.1

Horse Head to
Point of Shoals

Oyster reef
Sand-shell
Mud- shell
Sand
Mud

34.8
9.0

13.4

Point of Shoals

Dense

Scattering
Very Scattering
Depleted

26.9
13.1

5.5
2.0

Area III

Wreck Shoals

Dense

Scattering
Very Scattering
Depleted

48.6

Wreck Shoals to
Thomas Rock

Oyster reef
Sand-shell
Mud- shell
Sand
Mud

28.0

6.5

5.8

2.2

10.7

White Shoal

Dense
Scattering
Very Scattering
Depleted

57.0

10.3

9.1

Thomas Rock

Dense
Scattering
Very Scattering
Depleted

32.9
33.1
22.4
15.4

Further inspection of Moore's data reveals that the
present productive areas in the James River are in the
same approximate area as they were in 1910; however,
the areas of productive and potentially productive bottoms
may have increased since 1910. To show this, we compared
the geometric area of the top four categories shown by
Moore (Table 7) with our mud-shell, sand-shell and oyster
reef categories in Areas II and III. These data showed a
total area of 2,722 ha (6,727 acres) in 1910 and 4,534 ha
(11,204 acres) in 1980, a gain of about 60%. While this
cannot be considered conclusive because of the nature of

the original data set, the positive direction is suggestive. We
attribute the probable increase to the effect of culling
unwanted shells and small oysters onto unproductive sand
and mud bottoms from 1910 to 1981.

ACKNOWLEDGM ENTS

The authors thank P. Kendall, K. Walker and R. Morales-
Alamo for assistance in the field work; B. Bowen for his
contribution to statistical analysis; and the VIMS Art
Department for preparation of the figures.

Origin and Extent of Oyster Reefs in James River

151

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Natl. Shellfish. Assoc. 58:23-36.
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Va. Inst. Mar. Sci., Spec. Rep.. Appl. Mar. Sci. Ocean Eng.

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, D. S. Haven & D. B. Quayle. 1959. Fresh water kill of

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Bouma, A. H., ed. Shell Dredging and Its Influence in Gulf Coast

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Graves, C. 1905. Investigation for the promotion of the oyster

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247-341.
Haas, L. W. 1977. The effect of the spring-neap tidal cycle on the

vertical salinity structure of the James, York and Rappahannock

Rivers, Virginia, U.S.A. Estuarine Coastal Shelf Sci. 5:485-496.
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sp. (Haplosporida, Haplosporidiidae): causative agent of the

Delaware Bay oyster epizootic. Science 153 T414-1416.
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The effect of Tropical Storm Agnes on oysters, hard clams,

soft clams, and oyster drills in Virginia. Haven, D. S., W. J.

Hargis, Jr., J. Loesch, and J. P. Whitcomb, eds. Vie Effects of

Tropical Storm Agnes on the Oiesapeake Bay Estuarine Systems.

Baltimore, MD: John Hopkins Press. (CRC Publ. 54:488-407.)
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The use of sonic gear to chart locations of natural oyster bars in

lower Chesapeake Bay. Proc. Natl. Shellfish. Assoc. 69:11-14.
Haven, D. S., J. P. Whitcomb & P. Kendall. 1981a. The present and

potential productivity of the Baylor Grounds in Virginia. Va.

Inst. Mar. Sci, Spec. Rep. Appl. Mar. Sci. Ocean. Eng. No. 243:

1-154.
Haven, D. S., W. J. Hargis, Jr. & P. Kendall. 1981b. The oyster

industry of Virginia: Its status, problems and promise. 1981

(revised). Va. Inst. Mar. Sci., Spec. Pap. Mar. Sci. No. 4: 1024 p.
Hedgpeth, J. W. 1953. An introduction to the zoogeography of the

northwestern Gulf of Mexico with reference to invertebrate
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of the seed oyster reserves in Virginia and testing and identifying
gear to harvest oysters. Gloucester Point, VA: Va. Inst. Mar.
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133 p.

Loosanoff, V. L. 1931. Observation on Propagation of Oysters in
the James and Corrotoman Rivers and the Seaside of Virginia.
Newport News, VA : Virginia Comm. Fish. ; 46 p.

MacKenzie, C. L., Jr. 1983. To increase oyster production in north-
eastern United States. U.S. Natl. Mar. Fish. Serv. Mar. Fish. Rev.
45(3):l-23.

Marshall, N. 1954. Changes in the physiography of oyster bars in
James River, Virginia. Va. J. Sci. 5(3):23-28.

Moore, H. F. 1911. Condition and extent of oyster beds in the
James River. U.S. Bur. Fish. Doc. No. 729: 83 p.

Nichols, M. M. 1972a. Sediments in the James River, Va. Nelson,
B. W., ed. Environmental Framework of Coastal Plain Estuaries.
Geol. Soc. Am. Mem. 133:169-212.

. 1972b. Effect of increasing depth on salinity in the

James River estuary. Nelson, B. W., ed. Environmental Frame-
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571-589.

Nonis, R. M. 1953. Buried oyster reefs in some Texas Bays. /
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Pritchard, D. W. 1953. Distribution of oyster larvae in relation to
hydrographic conditions. Proc. Gulf Caribb. Fish. Inst.
5:123-132.

Scott, A. J. 1968. Environmental factors controlling oyster shell
deposits. Texas Coast. From Fourth Forum on Geology of
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Sokal, R. R. & F. J. Rohlf. 1981. Biometry, the Principles and
Practices of Statistics in Biological Research. San Francisco, CA:
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Stroup, E. D. & R. J. Lynn. 1963. Atlas of salinity and temperature
distributions in Chesapeake Bay 1952 - 1961 and seasonal
averages 1949-1961. Baltimore, MD:Chesapeake Bay Inst.,
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Journal of Shellfish Research, Vol. 3, No. 2, 153-167, 1983.

GENETIC DIFFERENTIATION AND POPULATION STRUCTURE OF THE

AMERICAN OYSTER CRASSOSTREA VIRGINICA (GMELIN)

IN CHESAPEAKE BAY

NORMAN E. BUROKER 1

Bureau of Biological Research

Rutgers, The State University of New Jersey

Piscataway, New Jersey 08854

ABSTRACT Genetic variation and differentiation were studied among 10 oyster bars of the American oyster Crassostrea
virginica (Gmelin) in Chesapeake Bay. The observed heterozygosity ranged from 0.195 to 0.230 while the proportion of
polymorphic structural loci ranged from 0.483 to 0.552 among demes. The genetic similarities among oyster bars averaged
99% suggesting little genetic differentiation; however, F s j statistics revealed that 23 of 41 alleles were significantly different
among demes, suggesting spatial heterogeneity among oyster bars within Chesapeake Bay. Principle component and step-
wise multivariate discriminant analyses of the 28 most common alleles indicated that the 10 oyster bars could be partitioned
into four different latitudinal groups (e.g., subpopulations). The four subpopulations are probably maintained by a balance
between the migration of planktonic oyster larvae and the adaptation of genotypes to local environmental conditions.

KEY WORDS: allelic variation, Chesapeake Bay, Crassostrea virginica, multivariate analysis, oyster bars, protein electro-
phoresis, subpopulations

INTRODUCTION

The American oyster Crassostrea virginica (Gmelin) is
an oviparous, dioecious bivalve. Its planktonic larval stage
lasts from two to three weeks and provides ample oppor-
tunity for zygotic dispersion (Galtsoff 1964). Large popula-
tions of sedentary adults of C. virginica are present in
Chesapeake Bay. Consequently, the possibility of genetic
differentiation between populations by genetic drift can be
discounted. In this study, I have attempted to delineate
between two theories concerning population structure of
C. virginica in the bay. The water circulation patterns
within the bay and the long planktonic larval stage provide
the potential for extensive gene flow among contiguous
oyster demes. These factors should contribute to high levels
of genetic similarity among oyster bars throughout the
latitudinal 240-km range of C. virginica in Chesapeake Bay.
This would suggest that the bay contains a single panmictic
oyster population. Alternatively, if selection pressure was
great enough to minimize the effect of gene flow among
oyster demes or if larval dispersion was not as wide spread
as suggested, then geographic variation in allele frequencies
could occur among various regions within Chesapeake Bay.
This effect would produce subpopulations (i.e., random
mating within groups) of the oyster in the bay instead of a
single panmictic population.

Examination of the population structure can be
approached through biochemical genetic studies using
protein electrophoresis on natural populations which will
provide information on gene and genotypic frequencies of
structural loci (Powell 1975, Selander 1976). Such informa-
tion may provide evidence for selection in natural popula-

Present address: Department of Biochemistry, School of Medicine,
The Oregon Health Sciences University, Portland, Oregon 97201.

tions in the form of macrogeographical clines in gene
frequency (e.g., spatial changes of allele frequencies with
concomitant geographical variation [Koehn 1969, Powell
1971, Schopf and Gooch 1971]); microgeographical clines
in gene frequencies (e.g., changes in allele frequencies with
concomitant microgeographic gradients or with local
environmental heterogeneity [Balegot 1971, Hamrick and
Allard 1972, Koehn et al. 1973]); temporal clines in gene
frequency (i.e., a progressive change in allele frequencies
with year-class or increasing age [Fujino and Kang 1968,
Koehn et al. 1971 , 1976, 1980, Tinkle and Selander 1973]);
and among-locus discordance in patterns of geographical
variation (Williams et al. 1973, Christiansen and Frydenberg
1974). In this analysis, 32 structural loci were examined in
C. virginica with respect to levels of genetic variation and
genetic differentiation among oysters bars in Chesapeake
Bay. The genie variation was then examined with respect to
environmental and geographical variations in the bay, and
to among-locus discordance between sampling localities.

Environmental Variation in Chesapeake Bay

Oceanic water moves along the lower water column of
Chesapeake Bay in a northerly direction to the head of the
bay. Fresh water enters the bay from tributaries and flows
in a southerly direction along the upper water column. This
opposite flow of fresh- and salt water at different depths in
the water column results in macro- and microgeographical
salinity gradients. High salinities are found in the deep
water layers and lower regions of the bay while lower
salinities are found in the surface water layers and upper
regions of the bay (Whaley and Hopkins 1952, Stroup and
Lynn 1963). This water circulation pattern may be the
reason why there are no large spatial gradients in water
temperature. Chesapeake Bay does experience seasonal

153

154

BUROKER

climatic variation. Winter surface water often freezes while
summer surface water temperatures may reach 30 C
simultaneously for all regions of the bay.

MATERIALS AND METHODS

Oyster Bars and Geographical Variation

The oyster samples used in this study were dredged from
ten oyster bars from various depths and regions of Chesa-
peake Bay and its tributaries (Figure 1, Table 1). Only
adult oysters of > 6 cm length were used. Individuals
smaller than 6 cm were not analyzed because of the possi-
bility of genotypic and age (size)-dependent interactions
which have been reported between marine bivalves (Koehn
et al. 1973, Mitton et al. 1973. Boyer 1974, Tracey et al.
1975, Singh and Zouros 1978, Zouros et al. 1980). All
samples were transported to the Marine Products Laboratory,
Center for Environmental and Estuarine Studies, University
of Maryland, where they were stored at — 20°C until
analyzed by starch gel electrophoresis. Both recruitment of
new individuals and ambient water conditions varied
between the collecting localities.

The spatial distribution of natural oyster bars in Chesa-
peake Bay ranges from the Swan Point site (upper bay) to
the James River (lower bay). This constitutes a latitudinal
geographic distance of approximately 240 km. The mean
water depths at the ten oyster bars sampled in this study
ranged from 0.3 to 6.1 m. A salinity gradient also exists
among the ten collecting localities, ranging from a mean
of 8.5 ppt at the Swan Point site to 17.5 ppt in Pocomoke
Sound. A very slight thermal cline in the annual mean water
temperature appears among the oyster bars which ranges
from 13.0°C at the Swan Point site to 15.5°C in the James
River (Table 1 ).

Sample Preparation, Electrophoresis and Protein Staining Systems

Approximately 0.5 g of either adductor muscle or
stomach tissue was extracted from each individual, placed
into a test tube containing 1.0 m2 of distilled water and
homogenized with a glass rod. This crude extract was
centrifuged at 5,000 rpm for 2 min. The supernatant was
then absorbed on cellulose wicks which were set into a
starch gel matrix. The methods of horizontal starch gel
electrophoresis of oyster samples including buffer solutions
and staining solutions are as previously described (Buroker
et al. 1975, 1979a, b). the 21 protein-staining systems used
in this study were: acid phosphatase (AcP), adenine kinase
(Adk), aldolase (Aid), aminopeptidase (Ap), aspartate amino-
transferase (Aat), esterase (Est), alphaglycerophosphate
dehydrogenase (aGlypd), glyceraldehyde 3-phosphate
dehydrogenase (Gly3pd), hexokinase (Hk), isocitrate
dehydrogenase (Idh), leucine aminopeptidase (Lap), malate
dehydrogenase (Mdh), malic enzyme (Me), mannose phos-
phate isomerase (Mpi), muscle protein (Mp), 6-phosphoglu-
conate dehydrogenase (6Pgd), phosphoglucose isomerase

(Pgi), phosphoglucomutase (Pgm), sorbitol dehydrogenase
(Sdh), tetrazolium oxidase ( To), and xanthine dehydro-
genase (Xdh). In this study the electrophoretic analyses of
soluble proteins reflected 32 structural loci. These loci were
selected on the basis of available staining procedures and
clarity of protein banding. Two polymorphic loci (AcP— 3
and Sdh) could not be resolved for all collecting localities.

Statistical Analyses

The inbreeding coefficient is the correlation between
random gametes within subdivisions relative to gametes of
the total population and is a measure of the heterogeneity
among the subpopulations (Wright 1940, 1969, 1978). The
variation in allele frequency between subpopulations can be
used to compute the "effective" inbreeding coefficient (F st ).
The estimate is

apj/p (1-p)

where p represents the weighted mean, and ah the weighted
sum of the squared deviations of the individual subpopula-
tions gene frequencies from the mean gene frequency divided
by the number of subpopulations:

'Pi =

= 2 [(p-pi) 2 /n]

Because each allele at a locus has its own values of ah and
p, F st can be used to test for differential selection between
the subpopulations. This statistic has also become widely
known and is used as the standardized variance of gene
frequency between populations (Cavalli-Sforza 1966).

Two other statistical procedures employed were principle
component and discriminant analyses. Principle component
analysis is a method of reducing the number of correlated
measurement variables into a small set of statistically inde-
pendent linear combinations having certain unique properties
with regard to characterizing individual differences (Overal
and Klett 1972, Harris 1975). The method (BMDP-4M;
Dixon 1977) was used here to describe biological, environ-
mental and genetic differences among oyster bars in Chesa-
peake Bay. A stepwise multivariate discriminant analysis
(BMDP-7M; Dixon 1977) procedure was used to select
those characters which best discriminate oyster subpopula-
tions in the bay.

RESULTS

Genetic Variation among Oyster Bars

The 21 protein-staining systems allowed examination of
32 monomorphic and polymorphic structural loci. The 18
loci which displayed genie variation have been tabulated
with relation to the collecting localities (Table 2). The 14
loci for which no genie variation was found are: AcP-\,
Adk-2,Ald, Aat-\,Est-2, Glypd-2, Gly3pdh,Hk-\,Me,
Mp-\, Mp-2, 7b- 1, 7c>-2, and Xdh. A summary of the

Crassostrea virginica in Chesapeake Bay

155

Figure 1. Map of Chesapeake Bay depicting the ten oyster bars sampled in this study. Starting at the head of the bay, the oyster bars are
Swan Point (SP), Herring Bay (HB), Broad Creek (BC), Tred Avon River (TAR), Patuxent River (PaR), Wicomico River (WR), Potomac
River (PoR), Pocomoke Sound (PS). Rappahanock River (RR), and James River (JR).

156

BUROKER

TABLE 1.

Biological and ambient environmental parameters from ten oyster bars in various regions of Chesapeake Bay.

The oyster bars range in decreasing latitude from Swan Point, MD, near the head of Chesapeake Bay

to James River, VA, near the mouth of Chesapeake Bay.

Oyster Bars

Maryland

TAR

PaR

PoR

Virginia

Mean recruitment

(spat/bushel)
(1939-1975) Maryland 2
(1961-1975) Virginia 3

Mean salinity (ppt)
(annual range)

Water depth (m) s

Mean water temperature
<°C) 4
(annual range)

14.6

8.5

121.4

12.5

32.1

12.5

36.4

11.0

15.4

12.0

63.6

14.8

!35.5 53.9

12.8

17.5

73.3

13.3

184.3

10.1

(3-14) (8-18) (8-18) (4-16) (8-18) (10-20) (8-18) (14-23) (8-18) (7-22)

5.2

13.0
(1-27)

4.6

6.1

0.3

4.6

3.7

3.0

3.7

13.8 13.5 13.6 13.6

(2-28) (2-28) (2-28) (2-28)

13.5 13.9 14.3 15.3 15.5

(2-28) (2-28) (2-29) (3-28) (2-28)

'SP (Swan Point), BC (Broad Creek), TAR (Tred Avon River), HB (Herring Bay), PaR (Patuxent River), WR (Wicomico River), PoR (Potomac

River), PS (Pocomoke Sound), RR (Rappahanock River), and JR (James River).
2 Source: Meritt (1977)

Source: D. Haven, Virginia Inst. Mar. Sci., pers. coram.
4 Sources: Whaley and Hopkins (1952). Stroup and Lynn (1963).

Source: G. Krantz. Marine Science Laboratory. Crisfield, MD 21817, pers. comm.

TABLE 2.
Genie variation of the American oyster Crassostrea rirginica among oyster bars from Chesapeake Bay.

Allele 2
Locus (RM)

Chesapeake Bay Oyster Bar;

TAR

PaR

PoR

.4p-l n

180

184

182

180

178

182

184

182

103

0.017

0.000

0.006

0.011

0.000

0.038

100

0.283

0.397

0.379

0.428

0.399

0.346

0.341

0.359

0.451

0.228

0.201

0.143

0.217

0.208

0.148

0.176

0.158

0.198

0.278

0.272

0.247

0.231

0.261

0.197

0.258

0.297

0.212

0.198

0.189

0.130

0.176

0.137

0.094

0.185

0.209

0.148

0.201

0.082

0.006

0.000

0.055

0.110

0.000

0.006

0.027

0.038

0.071

0.033

0.844

0.739

0.758

0.846

0.700

0.753

0.703

0.714

0.891

0.703

0.119

0.040

0.024

0.126

0.010

0.039

-0.059

-0.037

0.178

-0.008

AcP-3 n

—

148

—

178

168

182

184

120

—

0.000

—

0.000

0.024

0.000

115

—

0.014

—

0.035

0.051

0.014

0.065

—

0.005

0.011

110

—

0.507

—

0.593

0.439

0.444

0.589

—

0.198

0.446

108

—

0.291

—

0.267

0.398

0.389

0.202

—

0.357

0.353

105

—

0.169

—

0.105

0.112

0.153

0.119

—

0.324

0.174

100

—

0.020

—

0.000

—

0.115

0.016

—

0.541

—

0.744

0.857

0.806

0.750

0.593

0.587

—

-0.142

—

0.317

0.350

0.576

0.265

—

-0.171

-0.091

Adk-l n

180

186

182

184

178

182

176

184

104

0.000

0.006

0.000

0.005

0.000

102

0.078

0.059

0.088

0.103

0.124

0.067

0.088

0.108

0.082

100

0.233

0.226

0.198

0.239

0.191

0.315

0.258

0.193

0.196

0.130

0.572

0.597

0.665

0.582

0.640

0.522

0.582

0.625

0.603

0.592

0.117

0.118

0.049

0.076

0.045

0.090

0.071

0.074

0.114

0.185

0.000

0.011

0.600

0.548

0.593

0.674

0.551

0.674

0.637

0.580

0.630

0.652

0.002

-0.047

0.166

0.146

0.027

0.095

0.096

0.045

0.090

0.103

Crassostrea virginica in Chesapeake Bay 157

TABLE 2. Genie variation of the American oyster Crassostrea virginica among oyster bars from Chesapeake Bay (continued).

Locus

Allele 2

esapeaxe ba

y uyster ba

(RM)

TAR

PaR

PoR

176

186

180

112

178

166

114

117

0.000

0.006

0.000

109

0.006

0.000

0.006

0.000

0.006

0.012

0.009

100

0.761

0.715

0.761

0.773

0.778

0.789

0.723

0.787

0.747

0.711

0.233

0.285

0.228

0.227

0.222

0.211

0.277

0.208

0.241

0.281

0.364

0.462

0.400

0.364

0.267

0.374

0.446

0.337

0.410

0.456

-0.006

0.135

0.084

0.039

-0.228

0.133

0.116

-0.003

0.043

0.097

180

178

172

182

144

178

182

184

176

102

0.000

0.006

0.000

0.005

0.000

100

0.651

0.517

0.384

0.440

0.625

0.590

0.604

0.429

0.424

0.358

0.108

0.107

0.238

0.159

0.104

0.163

0.132

0.176

0.130

0.250

0.145

0.107

0.099

0.110

0.111

0.129

0.148

0.203

0.163

0.108

0.086

0.197

0.203

0.192

0.153

0.112

0.088

0.187

0.223

0.239

0.011

0.067

0.076

0.099

0.007

0.006

0.022

0.005

0.060

0.045

0.699

0.865

0.837

0.890

0.722

0.742

0.736

0.912

0.837

0.852

0.305

0.298

0.129

0.231

0.284

0.243

0.252

0.287

0.157

0.154

174

180

182

176

120

174

166

182

108

0.000

0.011

0.000

104

0.000

0.233

0.011

0.000

0.006

0.011

0.144

0.006

0.000

100

0.443

0.456

0.423

0.244

0.358

0.368

0.345

0.356

0.331

0.478

0.477

0.256

0.495

0.665

0.525

0.603

0.563

0.467

0.578

0.451

0.080

0.044

0.071

0.091

0.117

0.023

0.075

0.033

0.084

0.071

0.000

0.006

0.000

0.448

0.600

0.484

0.409

0.500

0.460

0.545

0.556

0.386

0.446

-0.204

-0.104

-0.154

-0.165

-0.143

-0.080

-0.022

-0.123

-0.297

-0.015

180

186

182

184

180

182

184

0.000

0.011

0.005

0.011

0.017

0.006

0.000

0.005

0.000

0.978

0.984

0.978

0.967

0.961

0.967

0.989

0.978

0.973

1.000

0.017

0.005

0.016

0.011

0.028

0.011

0.016

0.000

0.006

0.000

0.011

0.005

0.011

0.000

0.011

0.005

0.000

0.044

0.032

0.044

0.065

0.078

0.067

0.022

0.044

0.054

0.000

0.018

0.012

0.015

0.023

0.027

0.030

0.010

0.017

0.020

0.000

180

186

182

184

180

182

184

100

0.000

0.005

0.000

0.005

1.000

0.995

0.989

0.994

1.000

0.989

0.995

0.957

0.000

0.011

0.006

0.000

0.011

0.005

0.038

0.000

0.011

0.022

0.011

0.000

0.022

0.011

0.087

0.000

0.005

0.011

0.010

0.000

0.010

0.005

0.040

180

182

184

174

164

174

182

184

104

0.050

0.088

0.022

0.033

0.052

0.030

0.029

0.011

0.027

102

0.528

0.648

0.538

0.592

0.649

0.512

0.523

0.549

0.533

0.522

100

0.272

0.181

0.308

0.266

0.195

0.348

0.305

0.269

0.288

0.272

0.100

0.071

0.115

0.098

0.092

0.098

0.148

0.125

0.152

0.044

0.011

0.012

0.046

0.022

0.027

0.006

0.000

0.005

0.000

0.589

0.429

0.648

0.598

0.425

0.573

0.690

0.484

0.620

0.685

-0.069

-0.198

0.079

0.052

-0.196

-0.054

0.109

-0.199

0.005

0.088

180

182

180

184

178

180

182

170

182

178

0.000

0.011

0.000

0.011

0.000

0.005

0.011

0.067

0.077

0.117

0.082

0.129

0.056

0.066

0.071

0.096

0.811

0.747

0.717

0.736

0.778

0.797

0.718

0.747

0.803

0.122

0.159

0.167

0.185

0.124

0.167

0.137

0.206

0.176

0.090

0.000

0.005

0.000

0.005

0.000

0.006

0.000

0.311

0.319

0.411

0.370

0.360

0.333

0.385

0.329

0.407

0.348

-0.034

-0.223

-0.077

-0.169

-0.156

-0.085

0.125

-0.247

0.003

0.033

Aat-2

Est-l

Est-3

Idh-l

Idh-2

Lap -I

Lap-7

158 BUROKER

TABLE 2. Genie variation of the American oyster Crassostrea virginica among oyster bars from Chesapeake Bay (continued).

Allele 2
Locus (RM)

Chesapeake Bay Oyster Bars 1

TAR

PaR

PoR

Mdh-1 n

180

186

182

184

180

182

184

104

0.000

0.005

0.000

100

1.000

0.995

0.989

0.983

0.995

1.000

0.989

0.995

0.000

0.005

0.011

0.017

0.000

0.005

0.000

0.005

0.000

0.011

0.022

0.033

0.011

0.000

0.022

0.011

0.000

0.005

0.010

0.017

0.010

0.000

0.009

0.005

Mdh-2 n

180

186

182

184

180

182

184

103

0.006

0.005

0.000

0.006

0.000

0.005

0.011

0.005

0.000

0.994

0.984

0.989

0.995

0.989

0.983

0.984

0.989

0.995

0.000

0.005

0.011

0.005

0.006

0.017

0.011

0.005

0.000

0.005

0.000

0.011

0.032

0.022

0.011

0.022

0.033

0.022

0.011

0.010

0.008

0.010

0.005

0.010

0.017

0.014

0.013

0.009

0.005

Mpi-2 n

156

176

150

162

176

182

170

174

176

0.204

0.038

0.102

0.040

0.253

0.045

0.027

0.018

0.040

0.023

0.194

0.096

0.398

0.440

0.272

0.227

0.330

0.259

0.374

0.392

0.490

0.462

0.341

0.333

0.352

0.477

0.484

0.518

0.408

0.369

0.112

0.308

0.108

0.173

0.093

0.148

0.132

0.147

0.132

0.165

0.000

0.096

0.045

0.013

0.031

0.074

0.027

0.024

0.046

0.034

0.000

0.006

0.000

0.028

0.000

0.035

0.000

0.017

0.778

0.821

0.852

0.893

0.815

0.705

0.725

0.741

0.828

0.852

0.162

0.221

0.216

0.345

0.119

0.020

0.136

0.156

0.231

0.252

6Pgdh n

180

186

182

184

178

180

182

184

178

109

0.000

0.011

106

0.022

0.027

0.016

0.011

0.051

0.017

0.000

0.011

0.016

0.022

103

0.872

0.844

0.951

0.864

0.876

0.889

0.885

0.962

0.891

0.916

100

0.106

0.129

0.033

0.125

0.073

0.094

0.115

0.027

0.092

0.051

0.211

0.269

0.077

0.228

0.191

0.200

0.165

0.055

0.217

0.157

-0.073

-0.004

-0.186

-0.041

-0.146

-0.006

-0.194

-0.265

0.105

-0.007

Pgi n

180

186

182

178

180

182

184

114

0.006

0.000

0.005

0.006

0.000

0.005

0.000

110

0.022

0.044

0.055

0.028

0.039

0.049

0.044

0.054

0.016

106

0.672

0.747

0.703

0.643

0.702

0.656

0.659

0.703

0.674

0.739

100

0.289

0.210

0.242

0.297

0.258

0.300

0.286

0.236

0.261

0.239

0.011

0.022

0.005

0.000

0.006

0.005

0.011

0.005

0.000

0.005

0.000

0.444

0.387

0.505

0.473

0.449

0.467

0.484

0.473

0.522

0.370

-0.043

-0.024

-0.089

-0.047

0.023

-0.026

0.005

0.057

0.098

-0.066

Pgm-1 n

178

180

182

184

180

182

184

108

0.011

0.000

106

0.034

0.056

0.022

0.005

0.022

0.039

0.049

0.038

0.049

0.038

104

0.320

0.289

0.198

0.255

0.250

0.328

0.253

0.209

0.152

0.234

102

0.500

0.567

0.676

0.630

0.644

0.550

0.593

0.698

0.696

0.620

100

0.107

0.061

0.093

0.087

0.067

0.072

0.099

0.049

0.071

0.087

0.028

0.011

0.022

0.017

0.011

0.005

0.033

0.022

0.674

0.633

0.462

0.543

0.444

0.600

0.582

0.484

0.522

0.478

0.064

0.078

-0.067

0.027

-0.140

0.029

0.013

0.038

0.076

-0.134

Pgm-2 n

180

178

184

180

182

184

104

0.083

0.152

0.044

0.043

0.194

0.050

0.093

0.022

0.071

0.043

100

0.883

0.815

0.944

0.886

0.772

0.811

0.824

0.933

0.875

0.924

0.033

0.034

0.011

0.071

0.033

0.139

0.082

0.044

0.054

0.033

0.200

0.281

0.111

0.228

0.344

0.311

0.308

0.133

0.250

0.130

-0.048

-0.101

0.042

0.094

-0.055

-0.028

0.007

0.053

0.106

-0.091

Crassostrea virginica in Chesapeakf Bay

159

TABLE 2. Genie variation of the American oyster Crassostrea virginica among oyster bars from Chesapeake Bay (concluded).

Locus

Allele'
(RM)

Chesapeake Bay Oyster Bars

TAR

lilt

PaR

PoR

Sdh n

110
105
100

Number of loci studied 29

Mean number of genes

sampled per locus 177 ±15

148

—

178

182

0.043

0.061

—

0.051

0.027

0.034

0.038

0.034

0.891

0.858

—

0.854

0.896

0.898

0.901

0.909

0.065

0.081

—

0.096

0.077

0.068

0.060

0.057

0.217

0.284

—

0.247

0.187

0.159

0.198

0.182

0.087

0.117

—

-0.048

-0.023

-0.157

0.084

0.081

Polymorphic
loci/population

Heterozygous loci per individual

Observed

Expected

181 ±11 175 ±23 175 ±24 169 ± 23 173 ±26
0.483 0.483 0.517 0.552 0.552 0.552

176 ±17 171 ±28 182 ± 5 173 ±28

0.517

0.55:

0.517

0.214

0.216

0.215

0.230

0.195

0.216

0.222

0.196

0.227

0.218

±0.006

0.209

0.212

0.204

0.209

0.202

0.209

0.210

0.194

0.211

0.207

±0.006

'Oyster bars: Swan Point (SP), Herring Bay (HB), Broad Creek (BC), Tred Avon River (TAR), Patuxent River (PaR). Wicomico River (WR),
Potomac River (PoR), Pocomoke Sound (PS). Rappahanock River (RR), and James River (JR).

'Allele:

RM = relative allelic mobility on the gel
n = number of genes sampled per locus
H = observed heterozygosity

D = l(H

H e )/H e ]

where H n is the observed number of heterozygotes and H e is the Hardy-Weinberg expected

number of heterozygotes.

Based on 29 structural loci 'AcP— \,AcP— 3, and Sdh could not be resolved for all collecting localities).

genetic variation from the ten naturally occurring oyster
bars (Table 2) indicates that the proportion of polymorphic
loci (i.e., an estimate of the number of loci which exhibit
genie variation with the common allele [p] < 0.99 from a
random sample of structural loci in the population) ranges
from 0.483 to 0.552 among the ten collecting localities.
The observed heterozygosity (i.e., the proportion of genie
variation per individual) ranged from 0.195 to 0.230 among
the ten denies while the expected heterozygosities ranged
from 0.194 to 0.211.

In Table 2, the allele frequencies for the polymorphic
loci have been tabulated with respect to the geographic
spatial distribution of the oyster bars. No macrogeographic
cline is evident in any allele frequency across sampling
localities over the latitudinal distribution of Crassostrea
virginica in Chesapeake Bay. Also, little genetic differenti-
ation exists among the ten sampling localities based upon
estimates of genetic similarities and distances (Nei 1972).
That is, the genetic similarity among oyster bars ranged
from 0.985 to 0.998 while the genetic distances ranged
from 0.002 to 0.015. A Pearson chi-square statistic was
used to test the null hypothesis that the allele frequencies
are homogeneous among all oyster bars. This chi-square
analysis of the inter-oyster bar allelic contingency revealed
heterogeneity for 12 of the 18 polymorphic loci which
indicated among-locus discordance between oyster bars

for two thirds of the polymorphic loci sampled in this
study (Table 3).

Environmental versus Genetic Variation among Oyster Bars

In spite of a moderate level of environmental variation
and a great diversity in strength of recruitment among oyster
bars (cf. Table 1 ), there appears to be little genetic differ-
entiation among oyster bars in Chesapeake Bay based on
estimates of genetic distance, similarity, and variation
(above; Table 2). Principle component and stepwise multi-
variate discriminant analyses were employed to identify
any correlation between environmental variation and genetic
differentiation. The variables used were water depth,
temperature, salinity, and recruitment from Table 1. and
deme genie polymorphism and observed individual heterozy-
gosity from Table 2. The results of three principle component
plots involving principle components one, two and three
indicated that the ten oyster bars were diffused throughout
each plot with no clustering of collected localities. When a
stepwise multivariate discriminant analysis was conducted
between those localities from the upper (Swan Point.
Herring Bay, Broad Creek, and Tred Avon River) and those
from lower Chesapeake Bay (Patuxent River, Potomac
River. Wicomico River, Pocomoke Sound, Rappahanock
River, and James River), no discriminating variables were
found.

160

BUROKER

TABLE 3.

Inter-oyster bar allelic contingency tests for 18 polymorphic

loci in the American oyster Crassostrea virginica in

Chesapeake Bay.

Number of

Locus

Alleles*

Chi- square

d.f.

Probability

Ap-1

59.327

<0.001

Adk-1

71.750

<0.001

Aat-2

6.353

>0.700

AcP-3

124.720

<0.001

Est-1

133.374

<0.001

Est- 3

89.699

<0.001

Idh-1

9.886

>0.300

Idh-2

31.449

<0.001

Lap-\

66.789

<0.001

Lap-2

29.456

<0.050

Mdh-l

10.179

>0.300

Mdh-2

3.328

>0.900

Mpi-2

257.071

<0.001

6Pgdh

44.815

<0.001

Pgi

16.078

> 0.500

Pgm-l

192.985

<0.001

Pgm-2

86.173

<0.001

Sdh

5.447

>0.900

Total

1238.879

330

<0.001

*Rare alleles have been pooled with the next most common allele
for statistical reasons.

F st Analysis of Allele Frequencies

Although there were no apparent correlations of environ-
mental or geographical variables with levels of genetic
variation, among-locus discordance existed with respect to
spatial variation among oyster bars as indicated by the
contingency chi-square analysis of the polymorphic loci.
This suggested several possibilities: (1 ) differential selection
pressure for some alleles at these polymorphic loci in
response to the local environmental conditions of each
collecting locality, (2) a hierarchical relationship among
oyster bars, or (3) structuring of the dispersal pattern of
planktonic oyster larvae among the collecting localities in
Chesapeake Bay. A widely used method of revealing genie
heterogeneity between sampling areas is the use of the
standardized variance in allele frequencies, F st (Wright
1940, 1969. 1978; Cavalli-Sforza 1966; Neel and Ward
1972). F st estimates tend to be uniform for different
alleles when inbreeding, sample variation (genetic drift),
and random migration are occurring within a species. Con-
versely, natural selection operating independently on each
allele at a locus could reflect a heterogeneous array of F st
values among different alleles. That is, for alleles under
differential selection, the variance in allele frequency, as
well as the F st estimate, would be large. If there is a
balancing selection, the variance in allele frequency among
collecting localities would be small, as would the F st
values. A heterogeneity in F st values can also occur when a

hierarchical relationship exists among collecting localities,
or when migration is nonrandom and displays some pattern.

Table 4 gives the F st values for 41 alleles together with
the number of oyster demes over which they have been
estimated. Any allele with a mean allele frequency of (p) >
0.05 among oyster bars was analyzed. These data illustrated
a diversity of F st values for these alleles. The mean F st
value was 0.0161 and the variance Sp was 0.000194.

The F st statistic is related to the contingency chi-square
statistic used to test for heterogeneity between demes in
allele frequency estimates. Following the procedures of
Snedecor and Irwin (1933). this relationship can be
expressed as

< 2 = 2NF

st-

where N is the total sample size over all collecting localities.
In these circumstances F st is a simple function of the chi-
square statistic, with the significance of the chi-square
statistic. Consequently, when the chi-square values were
determined for the 41 F st values, 23 alleles were found to
be statistically significant, indicating heterogeneity for
more than one half of the alleles studied among oyster bars
in Chesapeake Bay (Table 4).

A further examination was made by F s t statistics in
testing for spatial heterogeneity among the oyster bars. The
test involved the goodness of fit of the observed distribu-
tion of F s t values to a theoretical chi-square distribution
(based on ten oyster bars) with 9 degrees of freedom
(Lewontin and Krakauer 1973). Table 5 compares the
observed distribution of the 41 F s j values in Table 4 with
chi-square distributions having 1, 2, 3, 4 and 9 degrees of
freedom. Classes were constructed to be one half of the
standard deviation of the mean F s t value. The theoretical
chi-square distribution with 9 degrees of freedom centers
on the observed mean F s t of 0.0161, while the other chi-
square distributions with 1 to 4 degrees of freedom were
not altered. The test for the goodness of fit between the
observed and the theoretical distributions gave a x 2 = 27.22
with 8 degrees of freedom corresponding to P < 0.001,
so the observed and theoretical chi-square distributions are
significantly different. A test for the goodness of fit between
the observed F s t distribution and a theoretical F s t distribu-
tion with fewer degrees of freedom provides better results
(Table 5). For example, the best fit occurs when 3 degrees
of freedom are selected as a mean for the theoretical chi-
square distribution. This indicates that the observed F s t
distribution of the 41 alleles compared among ten oyster
bars can best be explained if the ten oyster bars were con-
tained within four subpopulations from Chesapeake Bay.

Another test of heterogeneity is the comparison of the
observed variance of F s t with the theoretical variance as
described by Lewontin and Krakauer (1973). The theoretical
variance of F s t is given by the expression

Crassostrea virginica in Chesapeake Bay

161

TABLE 4.

F s , values for different allelic distributions among ten oyster
populations of Crassostrea virginica from Chesapeake Bay.

Locus Allele n*

st'

Chi-squareJ Probability Significant

TABLE 5.

Comparison of the observed distribution of F st values of

Table 4 with chi-square expected distributions having

means of 1, 2, 3, 4, and 9 degrees of freedom.

Expected

Ap-\

Acp-3

Adk- 1

Aat-2
Est-l

Est-3

Idh-i

Lap-l

Lap -2

Mpi-1

6Pgdh

Pgi

Pgm-1

Pgm-2

Sdh

100

97
94

100
96

102

100

96
94
92

92
88
84

103
100

106
100

104
102
100

104

100

110
105
100

10 0.00869
0.00635
0.00581

15.781
11.532
10.551

>0.05
>0.20
>0.20

No
No

Degrees of Freedom

Observed

0.01340

24.334

<0.01

Yes

< 0.0070

21.76

16.26

10.95

6.56

5.06

110

0.06102

62.851

<0.001

Yes

0.0071 - 0.0140

9.35

9.86

9.39

7.96

7.68

108

0.02095

21.579

<0.01

Yes

0.0141 - 0.0210

4.59

5.97

6.97

7.25

8.40

105

0.03449

35.525

<0.001

Yes

0.0211 - 0.0280

2.43

3.62

4.88

5.85

7.11

100
98
96

0.01275
0.00583
0.01813

23.129
10.576
32.888

<0.05
>0.30
<0.001

Yes
No
Yes

0.0281 - 0.0350
0.0351 - 0.0420
0.0421 - 0.0490
0.0491 - 0.0560

1.35
0.74
0.45
0.29

2.20
1.34
0.80
0.48

3.28

2.21
1.44
0.94

4.45
3.24
2.28
1.59

5.12
3.31
1.98
1.12

100

0.00421

6.568

>0.50

0.0561 - 0.0630

0.04

0.49

0.94

1.80

1.21

0.00421

6.568

>0.50

Total

41.00

100

0.04227

74.311

<0.001

Yes

x 2

11.03

6.26

5.95

17.87

27.22

98
96
94

0.01864
0.00837
0.01973

32.769
14.714
31.923

<0.001

>0.05

<0.001

Yes
No
Yes

df
Probability

5
>0.05

6 7
>0.30 >0.50

8
<0.05

8
<0.01

10 0.01888
0.04459

96 10 0.00544

10 0.00980
0.01150
0.00596

10 0.00662
0.00676
0.00834

10 0.04963
0.01833
0.02547

10 0.01346
0.01594

10 0.00516
0.00430

0.01434
0.01628
0.00409

10 0.03436
0.02604
0.02233

7 0.00289
0.00401
0.00239

30.548
72.147

9.923

17.542
20.585
10.668

11.135
12.141
14.979

80.401
29.695
41.261

24.443
28.947

9.391

7.826

26.041
29.564

7.427

52.914
40.102
34.388

2.769
3.842
2.290

<0.001
<0.001

>0.30

<0.05
<0.02
>0.20

>0.20
>0.20
>0.05

<0.001
<0.001
<0.001

<0.01
<0.001

>0.30
>0.50

<0.01

<0.001

>0.50

<0.001
<0.001
<0.001

>0.95
>u.90
>0.98

Yes

Yes
No

No
No
No

Yes
Yes
Yes

Yes
Yes

No
No

Yes
Yes
No

Yes
Yes
Yes

No
No
No

variance Sp = 0.000194

*n = number of oyster bars for each F st .
t^st = "effective" inbreeding coefficient.
$X 2 = 2NF st with (n - 1) degrees of freedom.

*corrected for the observed mean F S {.

o- 2 =KF st /(n- 1),

where K = 2 for an underlying binomial distribution of p
(the relative allele frequency). When this formula is applied
to the data of Table 4.

<r = 0.000058.

Whether or not the observed variance Sp , = 0.000194 is

r st

significantly larger than the theoretical variance was tested
by the ratio Sp Jo 2 = 3.368 which is distributed as x 2 /df.

To compensate for the multiple allelic loci, it was necessary
to remove 1 degree of freedom for each multiple allelic loci
(Lewontin and Krakauer 1973). The number of degrees of
freedom then became 41 - 14 = 27 and the probability of
the x 2 /df ratio was P < 0.001 . which indicates a significant
difference. If the assumption is made that the observed F s t
distribution can be best explained by assuming four sub-
populations in Chesapeake Bay instead of the actual ten
oyster bars studied, the a 2 becomes 0.0001 73. Whether the
observed variance is significantly larger was again tested by
the ratio

S 2 Fst /o 2 = 1.121.

The probability of the x 2 /df ratio was p > 0.25, resulting
in no significant heterogeneity.

The Sp Jo 2 ratio can be inflated by higher migration

rates among certain groups of subpopulations, by mutation
and special patterns of migration (Nei and Maruyama 1975),
as well as by hierarchical relationships among populations

162

BUROKER

(Robertson 1975). It is likely that higher migration rates
exist among neighboring oyster bars than among distant
oyster bars. Consequently, the partitioning of the ten oyster
bars in this study into four subpopulations greatly reduces
theSp Jo 2 ratio. The difference between the Sp Jo 2 ratios,
when 9 and 3 degrees of freedom were used, indicated how
this ratio can be inflated if the assumption is made that the
ten collecting localities were contained within a single
panmictic oyster population instead of four panmictic
subpopulations.

The F s t analysis indicates that some genetic structure
existed among the resident oyster bars of Chesapeake Bay.
Although the F s t analysis indicated subpopulational struc-
ture, it did not assign the ten oyster bars to their respective
group nor did it indicate which structural loci were respon-
sible for the partitioning of the ten bars into four subpopu-
lations. A probable solution to these questions can be
obtained with the use of principle component and stepwise
multivariate discriminant analyses. Using principle

component analysis, the Chesapeake Bay oyster population
was considered as a set and the allele frequencies of the
polymorphic loci as the variables measured over the ten
sampling localities. Twenty-eight of the most common
alleles (i.e., p > 0.05) from the Ap-l, Adk-l, Aat-2,
Est- 1 , Est-3, Lap- 1 , Lap-2, Mpi-2, 6Pgdh, Pgi, Pgm- 1 ,
and Pgm-2 loci (Table 2) were used in the principle
component analysis. The AcP—3 and Sdh loci were not
used because they were not represented among all oyster
demes. Also, the Idh—1, Idh-2, Mdh-l, and Mdh-2 loci
were not used because these genes represented very little
genie variation among demes and would not be of much
diagnostic value. The results of this analysis are illustrated
in Figure 2, where principle component one is plotted
against principle component three. The ten collecting
localities can be grouped into four subpopulations as shown
by the convex contour lines. The first group from the upper
Chesapeake Bay contains the Broad Creek, Patuxent River,
and Herring Bay oyster bars. Proceeding down the bay, the

4.0

3.0

2.0

1 .0 •

0.0

1 .0

-2.0

■3.0

■4.0

-5.0

I I

-4.0 -3.5

■3.0

I
■2.5

2.0

1 .5

■1.0

-0.5

I
0.0

0.5

1 .0

1 .5

2.0

2.5

3.0

3.5

PRIN 3

Figure 2. A principle-component analysis involving 28 alleles across 10 oyster bars in Chesapeake Bay. The two-dimensional graph depicts
principle components one and three of the analysis. Four different groups from Chesapeake Bay can be recognized. Group one is located in
upper Chesapeake Bay and consists of the Broad Creek (BC), Herring Bay (HB). and Patuxent River (PaR) oyster bars. Group two consists
of Swan Point (SP), Wicomico River fWR), and Potomac River (PoR) oyster bars. Group three consists of the Tred Avon River (TAR),
Pocomoke Sound (PS), and Rappahanock River (RR) oyster bars. Group four would contain the James River (JR) oyster bar. The contour
lines are drawn as a visual aid.

CRASSOSTREA VIRGINIA IN CHESAPEAKE BAY

163

second group contains the Swan Point, Wicomico River,
and Potomac River oyster bars. The third group encom-
passes the Tred Avon River, Pocomoke Sound, and Rappa-
hanock River oyster bars. The James River collecting
locality appears to be independent of all other groups. The
plots of principle components one on two and two on three
provided similar results. The separation of the ten oyster
bars into four different groups within Chesapeake Bay
supports the F s t analysis; however, it does not define which
alleles of the 12 polymorphic loci are diagnostic in parti-
tioning the ten oyster bars into four subpopulations.

An examination of allelic frequencies for the 18 poly-
morphic loci indicates that no single locus is diagnostic for
partitioning the oyster bars into subpopulations. yet there
are discrete allele frequency differences among the ten
oyster bars. Using stepwise multivariate discriminant analysis.

the information at these loci was combined to maximize
the diagnostic powers. Figure 3 shows the genetic differenti-
ation among nine oyster bars based on the first two
canonical variables of the discriminant analysis. The James
River oyster bar was excluded from the analysis because it
could not be grouped by principle components with any
other oyster bar. The Est-l 100 , Lap-\ 102 , Pgi 106 , and
Pgm— l 104 allele frequencies were used in combination
by the analysis to partition the remaining oyster bars into
three subpopulations.

DISCUSSION

The long planktonic larval development of Crassostrea
virginica has apparently been beneficial for the longevity of
this oyster species because it is this stage of development
that provides the opportunity for demes to disperse their

CD
O

c
o
c
co
O

20.0 >

16.0 •

12.0

8.0

4.0

0.0

-4.0

-8.0

1 2.0

16.0 •

20.0

\™«y

i i i i i i i i ■ i i i

30.0 -25.0 -20.0 -15.0 -10.0 -5.0 0.0 5.0 10.0 15.0 20.0 25.0

Canonical Variable 1

Figure 3. Two-dimensional graph of first two canonical variables from a stepwise multivariant discriminate analysis of 28 alleles across ten
oyster bars in Chesapeake Bay. The analysis entered the Est-l x<x> , Lap-l 102 , Pgi 106 , and Pgm-l [M alleles in the production of the two
canonical variables. Nine oyster bars can be distinctly grouped into three subpopulations. Group one consists of Broad Creek (BC), Herring
Bay (HB), and Patuxent River (PaR) oyster bars. Group two consists of Swan Point (SP), Wicomico River (WR), and Potomac River (PoR)
oyster bars. Group three consists of Tred Avon River (TAR), Pocomoke Sound (PS), and Rappahanock River (RR) oyster bars. The contour
lines are drawn as a visual aid.

164

BUROKER

zygotes into contiguous populations. As a consequence of
this dispersing ability, the fossil record indicates that ancient
populations of C. virginica apparently had relocated along
the Atlantic coast with respect to changing environmental
conditions (Merrill et al. 1965). The apparently high levels
of gene flow caused by large demes, high individual
fecundities, and long planktonic development should be in
part responsible for maintaining the relatively high level
of genetic variation within this species (Table 2). These
levels of genetic variation coincide well with those found
in other invertebrates (Lewontin 1974, Powell 1975,
Selander 1976). The genetic similarities between oyster
bars in Chesapeake Bay are consistent with levels of genetic
similarity reported for other intraspecific studies of inverte-
brates (Ayala et al. 1974, 1975; Hedgecock et al. 1976;
Tracey et al. 1975) as well as for other geographical popula-
tions of C. virginica along the Atlantic coast (Buroker 1983).

In spite of local environmental differences between the
ten collecting localities (Table 1), there appears to be no
overall genetic differentiation for the 32 loci studied among
the oyster bars. When the genetic variation among oyster
bars was statistically analyzed, however, gene diversity was
found and some structure to the oyster demes in Chesapeake
Bay was noted. The use of Wright's (1940, 1969, 1978)
inbreeding coefficient and principle component analysis
indicated at least four subpopulations of oysters in the bay.
The principle component analysis of the allele frequency
data produced the best possible grouping arrangement of
the ten oyster bars that were examined in this study. When
a comparison was made between sampling sites (Figure 1)
and their grouping (Figure 2), it became apparent that the
common factor which united the oyster bars within a group
was their close geographical proximity. For example, the
northern most group in Chesapeake Bay consisted of oyster
bars within the geographical boundaries of Broad Creek,
Herring Bay, and Patuxent River. Moving in a southerly
direction, the second group consisted of oyster bars within
the boundaries of the Wicomico and Potomac rivers. The
third group contained those oyster bars that were geographi-
cally bounded by Pocomoke Sound and Rappahanock River.
The final group consisted of oyster bars in the lower part
of the bay including the James River as part of the subpopu-
lation. These four subpopulations are latitudinally distrib-
uted in Chesapeake Bay. This may indicate that gene flow
among oyster demes was localized within certain regions of
the bay.

At least two observations can be made from a compari-
son of Figures 1 and 2. First, two oyster bars obviously
appear out of place. Based on geographic location, the
Swan Point oyster bar (group two) and the Tred Avon
River oyster bar (group three) would be more appropriately
placed in group one. Second, why should subpopulations
exist in Chesapeake Bay when there is a long planktonic
stage of larval development for this species and good water

circulation in the estuary? Because gene flow between
demes, which are in close geographical proximity, appears
to be a prominent mechanism in accounting for the oyster
population structure in the bay, either random events or
selection could be invoked to explain the above two obser-
vations. Because numbers of adult oysters per deme may
run from hundreds of thousands to millions of individuals
(Galtsoff 1964), it is unlikely that random events would be
responsible for the genetic differentiation found between
the Tred Avon River and Broad Creek oyster bars as well as
the Swan Point oyster bar and those demes of group one.
The alternative hypothesis would be genotypic adaptation
of new recruits to the local environmental conditions of
each oyster bar. For example, from the stepwise multi-
variate discriminant analysis (Figure 3). it is evident that
allelic variance of £sf-l 100 , Lap-V 02 , Pgi i0 \ and
Pgm-l iw is minimial within each group and is greatest
between subpopulations. Evidence for microgeographical
selection of allozyme genotypes in varied environments has
been presented for some marine bivalves (Koehn and
Mitton 1972; Koehn et al. 1973. 1976, 1980; Levinton
1973; Boyer 1974). Consequently, the microgeographic
adaptation of genotypes among Tred Avon River oysters to
their ambient environment might concide with that found
for oysters within group three instead of group one. The
same argument would place the Swan Point bar in group two
instead of group one. Obviously this results in a migration-
selection model to explain the genetic differentiation found
between sampling localities. When the balance within the
model is heavily shifted in favor of selection of genotypes
to local environmental conditions, the structuring of sub-
populations would be negated (which appears to be the
situation for Tred Avon River and Swan Point oyster bars).
On a macrogeographical scale the migration-selection
model can be used to explain the latitudinal partitioning of
oyster subpopulations in Chesapeake Bay. If there was no
opposing evolutionary force to counterbalance the effect
of gene flow, the bay would consist of a single panmictic
population with no genetic differentiation of groups;
however, the F s t analysis (Table 4) verifies genetic differ-
entiation among sampling localities when 23 of 41 alleles
were found to display significant heterogeneity among the
ten oyster bars investigated. Because subpopulations are
present that consist of minor genetic differences as revealed
by statistical analysis of allele frequencies among oyster
demes, it is suggested that a macrogeographical selection
gradient occurs latitudinally in the bay. Although the exact
components of this gradient cannot be defined, attention
can be drawn to some environmental similarities that
coincide with the subpopulations. The two most obvious
environmental parameters are salinity and water tempera-
ture, because both form latitudinal clines within the bay
(see Materials and Methods). Also, there is a positive rela-
tionship between eigenvector coefficients of the principle

CRASSOSTREA VIRG1NICA IN CHESAPEAKE BAY

165

component analysis for oyster spat recruitment, salinity,
water depth of the oyster bars, deme genie polymorphism,
and observed individual heterozygosity (see Results). It
must be assumed that some environmental selective effect
existed that counteracted migration to establish a balance
among these evolutionary forces (i.e., migration and selec-
tion) and was responsible for the partitioning of subpopula-
tions. A possible candidate would be Haplosporidiwn
nelsoni (Haskin) (MSX) disease which has a history of
periodically reoccurring among oyster populations along
the Atlantic coast of North America (Andrews and Wood
1967. Haskin and Ford 1982). In Chesapeake Bay, this
disease has at times produced heavy mortality among
oyster bar in the high-salinity areas of the lower bay while
the disease has had no apparent affect among oysters that
inhabit the low-salinity environment of the upper bay.

The occurrence of bivalve subpopulations has been
hypothesized in some instances to explain heterozygote
deficiencies among allozyme genotypes of Mytilus
californianus (Conrad) (Tracey et al. 1975), Mytilus edulis
(Linnaeus) (Boyer 1974), Tridacna maxima (Roding)
(Ayala et al. 1973), and Crassostrea virginica (Zouros et al.
1980). If the allele frequencies from two genetically
different sampling localities are pooled and Hardy-Weinberg
equilibrium frequencies are estimated as if the sample
represented one population, the expected frequencies of
heterozygous individuals would be over estimated (i.e.,
the Wahlund effect). When considering the Wahlund effect,
it is important to emphasize that (1) the allele frequencies
among denies must be different to be able to detect a signi-
ficant deficit of heterozygous genotypes, and (2) the effect
is uniform over all loci which exhibit genie differentiation
among denies. A measure commonly used to detect hetero-
zygote deficiences among allozyme genotypes is the "D"
statistic (Koehn et al. 1973) in which a positive value
indicates a heterozygote excess and a negative value a
heterozygote deficit. This value has been recorded in
Table 2 for all polymorphic loci across the ten sampling
localities. A predominant deficit of heterozygous genotypes
was found for the Est-3, Lap-2, and 6Pgdh loci over the
ten sampling sites. Zouros et al. (1980) also reported hetero-
zygote deficits for Est— 3 and Lap-2 among their largest
weight classes of C. virginica collected from Malpeque Bay,
Nova Scotia. Canada. Contrary to the deficiency of hetero-
zygotes. an excess of heterozygote genotypes occurred at
the Adk—l, Est— I, and Mpi-2 loci across the ten oyster
bars. Of these six loci, significant heterogeneity occurred
across oyster bars (ref: F s t analysis. Table 4)forthe.4cr7c— 1 ,
Est— 1, Est -3, Mpi-2, and 6Pgdh loci. The Wahlund effect
may be responsible for the deficit of allozyme heterozygotes

which has been reported among marine bivalves if only two
alleles are involved. When three or more alleles are present,
it is possible to generate an overall excess of heterozygous
genotypes (Li 1969, Milkman 1975, Koehn et al. 1976).
The Est-3, Lap-2, and 6Pgdh loci generally have two
common alleles while the Adk—l, Est-l, and Mpi— 2 loci
have three or more common alleles. Consequently, the
Wahlund effect can explain both excesses and deficits of
heterozygous genotypes within a collecting locality.
Balancing selection has also been used to explain hetero-
zygote deficits in marine bivalves (Koehn and Mitton 1972;
Koehn et al. 1973, 1976; Mitton et al. 1973; Boyer 1974)
while a heterozygote advantage has been used to explain
heterozygote excesses among bivalves (Buroker 1979.
Fujioetal. 1979, Zouros et al. 1980).

In conclusion, the levels of genetic variation for
Crassostrea virginica in Chesapeake Bay coincide well with
those found for other geographical populations along the
Atlantic coast of North America (Buroker 1983). Although
the genetic distances among oyster bars in the bay were
small, interdemic heterogeneity was found for 23 of 41
alleles tested. It was by means of this among-locus variation
across oyster bars that subpopulations were classified
primarily by principle component and discriminant analyses.
The subpopulations and among-locus variation across oyster
bars are thought to be maintained through a balance
between migration and selection. Because this study only
draws attention to the possibility of population differenti-
ation among marine bivalves with long planktonic stages of
larval development, the findings of this report should be
thoroughly tested by investigations which analyze the
temporal genetic stability of recruits as compared to
resident individuals.

ACKNOWLEDGMENTS

I express my gratitude to the Maryland Sea Grant
Program (under the National Oceanic and Atmopsheric
Administration, United States Department of Commerce);
the Marine Products Laboratory, Center for Environmental
and Estuarine Studies, University of Maryland; and the
Charles and Johanna Busch endowment. Bureau of Biological
Research. Rutgers University, which have in part supported
this research. I also thank Michael E. Douglas. Richard K.
Koehn, James F. Leslie, Roger D. Milkman, and Robert C.
Vrijenhoek for their constructive criticism; and William
Browning, Evelyn Dicey, and Diane Pruitt for their assistance
throughout this study. Very special thanks are accorded to
George Krantz for the oyster samples used in this study and
for providing information on the history of the Chesapeake
Bay oyster bars.

166

BUROKER

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Journal of Shell fish Research. Vol. 3, No. 2, 169-174, 1983.

FEASIBILITY OF MARICULTURE OF THE HARD CLAM

MERCENARIA MERCENARIA (LINNE)

IN COASTAL GEORGIA

RANDAL L. WALKER

Skidaway Institute of Oceanography
P.O. Box 13687
Savannah, Georgia 31416

ABSTRACT Caging, baffles, and the utilization of feeder creeks and tent structures were tested as predator controls to
determine if mariculture of the hard clam Mercenaria mercenaria (Linne) is feasible in coastal waters of Georgia. Clams
were planted at densities of 509, 1009, 2018, and 3027 clams m~ 2 in replicate plots within predator-free cages on an
intertidal sandflat at Cabbage Island, Wassaw Sound, Georgia. Seed clams grew from a mean shell length (SL) of 6 to 24 mm
in 7 months with no observed density effects on survival or growth (ANOVA a = 0.05). Seed clams (4,000 of 10-mm SL
and 40 of 25-mm SL) were planted in replicate plots (2.25 m 2 ) of baffles and mud, baffles, mud and tops, baffles and
gravel, and baffles and mixed oyster shells. No differences in survival were evident between test treatments for either seed
size (ANOVA °= = 0.05). Seed clams (76 of 30-mm SL and 4,000 of 5-mm SL) were planted on the bottom of a feeder
creek in replicate plots of mud, mud plus mixed oyster shells, mud plus tent, and mud with shells and tent. Survival for the
30-mm SL seed clams ranged from 43 to 86% and was < 1% in all plots for 5-mm SL seed clams. No differences in percent
survival were evident between plots for either size class (ANOVA cc= 0.05). A plan is presented for hard clam mariculture
in the coastal waters of Georgia.

KEY WORDS Hard clams, Mercenaria mercenaria, mariculture, predation. Georgia.

INTRODUCTION

The hard clam Mercenaria mercenaria (Linne) represents
a new and potentially important supplemental fishery for
the state of Georgia. The potential of the fishery is viewed
primarily as an off-season fishery for the blue crab
(Callinectes sapidus [Rathbun] ) fishermen in Georgia.

The extent of the hard clam resource in Georgia is
unknown. Godwin (1967, 1968) conducted a clam survey
of 432 stations in inshore Georgia waters (estuaries and
creeks). Clams occurred at 41 stations, primarily in inter-
tidal, higher saline areas with sand-mud or sand-mud-shell
substrates. Godwin (1967) reported a maximum clam den-
sity of 16 m" 2 (151 • 100 ft" 2 ) with a mean density of
5 clams m~ 2 , and concluded that, at that time, a commercial
hard clam fishery was not feasible.

In more northern U.S. waters, M. mercenaria occurs pri-
marily subtidally in sounds or estuaries. In Georgia, however,
most clam beds in Wassaw Sound are located intertidally in
the headwaters of major creeks, in the small feeder creeks,
among live oysters, or among oyster-shell deposits associated
with live oyster reefs at densities of up to 100 clams m~ 2
(Walker et al. 1980, Walker and Tenore [in press]). Areas
with densities greater than 25 clams m~ 2 are not uncommon
in the small feeder creeks and in the headwaters of the major
creeks of the higher saline areas of the Sound. These
densities are much higher than those reported by Godwin
(1967) and could support a small commercial fishery.
Though dense, these clam beds are small and are easily
overfished. For example, one clam bed of approximately
90-m 2 area which had a mean density of 49 clams m" 2
occurs in a feeder creek (3X61 m) located on a Wassaw

Island, a National Wildlife Refuge. This bed was illegally
fished in 1981 and the mean density decreased from 49 to
21 clams m" 2 within a week (Walker, in preparation). Suscep-
tibility to overfishing may explain the sporadic nature of
the hard clam fishery during the past 103 years in Georgia
(Walker et al. 1980).

One method of maintaining clam production in Georgia
may be through clam mariculture. Clams grow year around
in the coastal waters of the southeastern United States
(Eldridge et al 1976), and Georgia has vast areas suitable
for shellfish culturing. The major problem with clam
mariculture in Georgia is predation by blue crabs (Walker
et al. 1980) and mud crabs (Panopeus herbstii [Say] )
(Whetstone and Eversole 1977). Methods for reducing clam
predation include fencing, caging, utilizing various types of
aggregates (gravels, shell), and baffles (Kraeuter and Castagna
1980, Castagna and Kraeuter 1981). The purpose of this
paper is to discuss the feasibility of clam mariculture in the
coastal waters of Georgia.

Study Site Location

Wassaw Sound (Figure 1 ) is an estuarine embayment
located in the Georgia Bight (Howard and Frey 1980). Tides
are semidiurnal and average 2.4 m amplitude, with spring
tides ranging approximately 3.4 m (Hubbard et al. 1979).
Water temperatures range from 8 to 30°C (Dorjes 1972)
and salinities at the mouth of the Sound range from 20 ppt
in the winter to 30 ppt in the summer (Howard and Frey
1980). Sediments range from silty clay to fine sand; inter-
bedded sand-mud is the most prevalent sediment (Howard
and Frey 1975).

169

170

Walker

Figure 1. Wassaw Sound showing various experimental study sites.

MATERIALS AND METHODS

Clams were obtained from Aquaculture Research Corpor-
ation, Dennis, Massachusetts, in June 1982, and planted
and maintained at different densities (509, 1009, 2018, and
3027 m' 2 ) within 1 - X 1 - X 0.3-m cages constructed of 3-mm
mesh Vexar® plastic. The plastic mesh was attached to
1- X 1-X 1-m frames constructed of 1 3-mm steel reinforcing
rods. The resulting structure was buried to a depth of
0.85 m into a sandy sediment on an intertidal flat at
Cabbage Island. G A (Figure 1 ). Cages were sampled monthly
to a sediment depth of 10 cm, with sediment, clams and
crabs sieved through a 5-mni mesh screen. Clams were
counted and a subsample (n = 70) was measured for shell
length (anterio-posterior measurement); the clams were
then returned to their original plot. Crabs were identified as
to species, measured for carapace width, and discarded.
Differences in clam growth rates were determined by
analysis of covariance (ANCOVA).

In October 1982, 2.25-m 2 replicate plots of baffles and
mud, baffles, mud and tops, baffles and 151 C of mixed
oyster shell, and baffles and 151 C of crushed aggregate
gravel (No. 57, coarse; mean diameter = 5 cm) were set up
on an intertidal mud flat at House Creek (Figure 1). Baffles
were constructed of 6-mm mesh Vexar® plastic attached to
a 13-mm steel reinforcement rod. Four baffles were buried
in the mud at right angles to each other until the Vexar®
plastic was 0.15 m deep. Baffles were allowed to stand
2 months before the shell, gravel, or tops were added. After
two weeks, 40 seed clams of 2-cm SL (18 m~ 2 ) and 400
seed clams of 1 -cm SL ( 1 78 m" 2 ), each size class identifiable
by color codes using Krylon® spray paint, were placed into
each plot. Tops constructed of 25-mm mesh Vexar®
plastic (i.e., bird netting) were attached to two plots after
seeding. Plots were sampled as above in July 1983. The
mean survival percentage was determined for each plot and
clam size class. The resulting data were examined statistically
by analysis of variance (ANOVA).

Feasibility of Mariculture of Hard Clam in Georgia

171

In October 1982. experimental plots (3.7 X 4.3 m) were
established in two mud-bottom feeder creeks located at
Wassaw Island (Figure 1). Wild clams were removed by
raking. The following replicate plot types were set up: mud
bottom, mud bottoms with 151 £ of washed oyster shell,
mud bottoms with a "tent structure," and mud bottoms
with 151 £ of shell and a tent structure. Tents were con-
structed of 13-mm Vexar® plastic which was cut into
3.7 X 4.3-m sections. Along the long sides 13-mm steel
reinforcement rods were attached and along the short
sides 3.7 m of 13-mm galvanized chain was attached and
buried in the substrate. Four, 15-cm diameter styrofoam,
crab-trap floats were attached along the midline of the
structure. At low tide the structure rested flat on the
bottom and floated into a pup-tent form as the tide entered
the creek.

Each plot was seeded with 4,000 clams of 0.5 cm SL
(251 m" 2 ) and 76 clams of 3.0-cm SL (5 irf 2 );each size
class was color coded using Krylon® spray paint. In
addition, four plots, one of each test variable, received 80
of 1-cm SL and 45 of 2-cm SL seed clams. Whole plots were
sampled in August by sieving sediments to a depth of 10 cm
and clams through a 5-mm mesh screen. The clams were
counted and shell lengths were determined using vernier
calipers. The numerical data were statistically analyzed by
analysis of variance.

RESULTS

No significant differences existed in growth rates
(ANCOVA cc = 0.05), in final shell length (ANOVA « =
0.05), or in survival per density per month (ANOVA oc =
0.05). Clams grew from an initial mean shell length of 6.1
to 23.9 mm (Table 1) from June 1982 to January 1983.
Growth rates at each density are given in Figure 2.
Instantaneous clam survival was lowest in July (76.5%)
but exceeded 99.0% by October (Table 2).

In the baffle experiment, total clam survival was greater
for the 2.5-cm seed clams (72%) than for the 1.0-cm clams
(23%). No significant differences existed between protec-
tive methods (ANOVA <* = 0.05) for either the 2.5- or 1 .0-cm
clams; however, survival percentages per treatment were as
follows: (1.0-cm size class) mud, baffles, and top (29%),
mud and baffles (26%), shell and baffles (23%), and gravel
and baffles ( 1 l%);(2.5-cm size class) mud and baffle (88%),
shell and baffle (73%), mud. baffle, and top (68%), and
gravel and baffle (45%).

In the tidal creek experiments, total clam survival
decreased with decrease in clam size as follows: 3 cm
(66%) > 2 cm (57%) > 1 cm (40%) > 0.5 cm (0.04%).
For the 3-cm size class, no significant differences existed
between plots (ANOVA « = 0.05); however, clam survivals
per treatment were as follows: shell and tent (86%) > mud
and tent (70%) > shell (62%) > mud (43%). For the 5-mm
clams, no significant differences existed in clam survival
between plots (ANOVA <* = 0.05). Mortality of 5-mm clams

exceeded 99% in all plots. For the 2-cm clams, 76% were
recovered from shell and tent plots, 64% from mud and
tent plots, 53% from shell plots, and 4% from mud plots.
For the 1.0 cm clams, 81% were recovered from the shell
and tent plot, 73% from the shell plot, 4% from the mud
plot and 0% from the mud and tent plot.

DISCUSSION

Mortality reduction is essential to any successful clam
mariculture program. The size(s) at which hard clams are
no longer preyed upon by different predators are as follows:
Cancer irroratus Say (1 5 mm)(MacKenzie 1977), Urosalpinx
cinerea Say (20 mm) (MacKenzie 1977), Panopeus herbstii
(35 mm) (Whetstone and Eversole 1981), Callinectes sapidus
(40 mm) (Arnold 1983). and Menippe mercenaria (Say)
(70 mm) (Arnold 1983). Whelks of the genus Busycon prey
on all sizes of clams (Peterson 1982). If unprotected seed
clams are planted in the field, they are preyed upon by a
host of predators. Menippe mercenaria and Busycon whelks
are capable of preying upon commercial-size clams. Thus,
some means of protection is mandatory.

Clam mariculture attempts in the field using various
methods of seed protection have had mixed success. Seed
clams (10 mm) which were planted in unfenced and fenced
plots with 1.8-m high and 13-mm mesh plastic screen in
Florida resulted in 100% mortality (Menzel and Sims 1964).
Clams ranging from 33- to 44-mm SL that were planted in
those plots suffered 100% mortality in the unfenced plots
and less than 5 to 18% mortality in fenced plots (Menzel
and Sims 1964). Seed clams that were planted in fenced
plots in Virginia averaged 94% survival as compared to 8.8%
for those in unfenced plots (Kraeuter and Castagna 1980).
Field experiments in Virginia, using crushed rocks, pea
gravel, crushed oyster shell, or whole oyster shell as pro-
tective cover for 0.6 to 20-mm seed clams, resulted in
survivals greater than 80% as compared to 15 to 35%
survival in unprotected control plots (Castagna 1970);
however, in Florida, the survival rate of seed clams (4 to
20 mm) was 10% in plots with pea gravel, 2% in crushed
oyster shell, and less than 1% in controls (Menzel et al.
1976). Fenced clam plots in Chesapeake Bay did not
increase survival; however, gravel did increase clam survival
by 10% for 2 to 17-mm seed clams (Haven and Loesch 1973).
Thus, depending upon location and environmental condi-
tions, the use of these protective measures may result in
widely varying rates of survival.

Acceptable survival in Georgia (> 70%) resulted for seed
clams greater than 20 mm when protected by baffles,
baffles and shell, shell and tents, and for seed clams greater
than 10 mm when planted in cages. The survival of seed
clams in cages is dependent upon the monthly removal of
crabs, Callinectes sapidus and/or Panopeus herbstii, which
probably entered cages as metamorphosing juveniles
(Figure 3). In similar cages, which were planted earlier at
the same site, seed-clam (10 mm) survival in the first year

172

Walker

TABLE 1.

Growth (in mm) of hard clams that were planted at densities of 509, 1009, 2018, and 3027 m
on an intertidal sandflat at Cabbage Island, Georgia.

Densities

509 m~ 2

1009 m 2

2018 m"

-2

3027 m

-2

Date

Plot 1

Plot 2

Plot 1

Plot 2

Plot 1

Plot 2

Plot 1

Plot 2

June 1982

6.1

July 1982

8.5

8.4

8.7

8.8

8.9

9.0

8.2

8.9

August 1982

11.0

12.1

11.9

12.0

12.7

11.3

12.3

11.1

September 1982

13.3

11.9

13.2

14.0

12.0

13.0

12.4

13.0

October 1982

18.3

16.4

17.8

18.0

16.9

17.9

17.5

17.1

November 1982

23.3

21.3

22.6

23.7

21.5

22.1

21.5

December 1982

24.0

21.4

22.7

24.4

21.5

23.3

22.2

21.5

January 1983

25.2

22.7

24.6

25.7

22.3

25.0

23.5

22.3

c
d>

|- j 500m" 2 y= 5.55x 071 r 2 0.9677
\ 1000m" 2 y= 5.63x 072 r 2 0.9777
{ 2000m" 2 y=5.70x 069 r 2 9715
[ 3000m"2 y = 5.67x 0B8 r 2 0.9741

r SjD.

Jun Jul Aug Sept Oct Nov Dec Jan
Time(months)

Figure 2. Growth (in mm) of hard clams, Mercenaria mercenaria,
planted at different densities in cages at Cabbage Island, GA. Data
points represent the mean of replicate densities. Y = shell length (in
mm) and x = time (in months).

TABLE 2.

Mean survival percentage per clam density per month for

hard clams that were planted in protective cages on an

intertidal sandflat at Cabbage Island, GA.

Densities

Overall

Date

509 2

1009 m~ 2

2018 m 2

3027 m 2

Survival

July 82

89.3

66.2

83.3

59.5

76.5

Aug 82

86.6

95.0

87.6

85.9

87.9

Sep 82

83.0

97.3

94.8

95.5

94.6

Oct 82

100.0

99.1

99.7

99.6

Nov 82

99.1

99.5

100.0

99.2

99.5

Dec 82

100.0

99.1

100.0

99.9

Jan 83

98.2

97.7

99.1

99.7

99.1

Final mean

survival

percentage

62.5

59.0

67.9

48.1

62.4

6 1

5 "

-Q

1 ■

Blue Crabs

I — I Mud Cr

abs

Aug Sep Oct

1982

Nov

Dec

Jan

1983

Figure 3. Number and species of crabs removed monthly from
experimental clam cages located on an intertidal sandflat at Cabbage
Island, GA. (The number in parenthesis is the mean carapace width
per crab species.)

ranged from 14 to 31% because crabs were not removed
monthly but seasonally and they grew large enough to prey
upon the clams within the cages (Walker, in preparation).

The following mariculture program is considered feasible
for the coastal waters of Georgia. Seed clams (6 mm),
which are planted at densities of up to 3,027 m" 2 , can be
grown to a shell length greater than 20 mm within 7 months
with greater than 80% survival if they are planted in spring/
summer and if crabs are removed from their cages at least
monthly. Once the clams reach a shell length of 25 mm,
they can be transplanted into plots with baffles or into
creeks using shell cover and/or tent structures as protective
cover, or left in cages after densities are reduced (Walker.
in press).

In Georgia, small feeder creeks (defined as being generally
less than 4.5 m in width and several hundred meters in length)

Feasibility of Mariculture of Hard Clams in Georgia

173

appear to be the best habitat for clam mariculture. Feeder
creeks generally drain at low tide or retain standing pools
behind oyster reef "dams" which are located at the mouth
of or within the creeks. Wild clams may occur in high
densities (up to 100 nf 2 ) within feeder creeks. The growth
rates of clams in feeder creeks do not differ from those
from other habitats; however, clams usually grow faster in
sandy substrates (Rhoads and Panella 1970, Kennish and
Olsson 1975).

Many clam predators in Georgia do not occur in feeder
creeks. The Atlantic oyster drill Urosalpinx cinerea (Say)
usually occurs at the mouth of and rarely within feeder
creeks (Walker 1981). The southern oyster drill Thais
haemastoma (Conrad) and the starfish Asterias forbesi
(Desor) have not been found in feeder creeks (Walker
1982). The whelks, Busycon carica (Gmelin) and B.
contrarium (Conrad) generally do not occur in feeder
creeks. These creeks also provide a physically less dynamic
environment than do major creeks or open areas of the
sounds which are exposed to wave action.

Baffles, cages and pens, which were placed in major
creeks or open areas of the sounds in Georgia, have not

been successful in protecting clams. Cages that were buried
in sandy sediment on intertidal flats at Cabbage Island and
anchored with approximately 16 kg of bricks were washed
out during winter storms, completely buried by shifting
sediments, or so severely damaged that clams washed out
(Walker, in preparation). Baffles, which were placed in
feeder creeks, creeks or areas of the open sound, have met
with similar fates. Furthermore, pens, cages and baffles were
often vandalized by boaters and sports fishermen or run over
by boaters. Beds that are located in small feeder creeks
are nominally protected from boats, people (vandals), and
wave action, and represent a valuable fishery resource to
the state of Georgia.

ACKNOWLEDGMENTS

The author thanks Drs. E. Chin, J. Harding, R. Mann,
D. Menzel, and K. R. Tenore plus two anonymous individ-
uals for reviewing the manuscript. Special thanks are given
to Ms. A. Boyette and Ms. Suzanne Mcintosh for the
graphics, and to Ms. L. Land for typing the manuscript. The
work was supported by the Georgia Sea Grant Program
under grant number USDC-RF/8310-21-RR100-102.

REFERENCES CITED

Arnold, W. S. 1983. The effects of prey size, predator size, and
sediment composition on the rate of predation of the blue crab
{Callinectes sapidus Rathbun), on the hard clam (Mercenaria
mercenaria Linne). Athens, GA: LIniv. of Georgia; Thesis. 47 p.

Castagna, M. A. 1970. Field experiments testing the use of aggregate
covers to protect juvenile clams. Proc. Natl. Shellfish. Assoc.
70:2 (Abstract).

& J. N. Krauetei. 1981. Manual for growing the hard

clam, Mercenaria. Va. Inst. Mar. Sci. Spec. Sci. Rep. Appl. Mar.
Sci. Ocean Engin. No. 249; 110 p.

Dorjes. J. 1972. Georgia coastal region, Sapelo Island, U.S.A.:
Sedim'entology and biology. VII. Distribution and zonation of
macro-benthic animals. Senckenb. Marit. 4:182-216.

Eldridge, P. J., W. Waltz, R. C. Gracy & H. H. Hunt. 1976. Growth
and mortality rates of hatchery seed clams, Mercenaria mercenaria
in protected trays in waters of South Carolina. Proc. Natl.
Shellfish. Assoc. 66:13-20.

Godwin, W. F. 1967. Preliminary survey of a potential hard clam
fishery. Ga. Game Fish Comm. Contrib. Ser. 1:23 p.

. 1968. The distribution and density of the hard clam,

Mercenaria mercenaria, on the Georgia coast. Ga. Game Fish
Comm. Contrib. Ser. 10:30 p.

Haven, D. S. & J. G. Loesch. 1973. An investigation into commer-
cial aspects of the hard clam fishery and development of com-
mercial gear for the harvest of molluscs. Gloucester Point, VA:
Virginia Institute for Marine Science; Annual Contract Rep.
No. 3-124F;91 p.

Howard, J. D. & R. W. Frey. 1975. Estuaries of the Georgia coast,
U.S.A.: Sedimentology and biology. II. Regional animal-sediment
characteristics of Georgia estuaries. Senckenb. Marit. 7:33-103.

. 1980. Holocene depositional environments of the Georgia

coast and continental shelf. Howard, J. D.. C. B. DePratter and
R. Frey, eds. Excursions in Southeastern Geology: Tlie
Archaeology-Geology of the Georgia Coast. Guidebook 20:
66-134.

Hubbard, D. K., G. Oertel and E. Nummendal. 1979. The role of

waves and tidal currents in the development of tidal-inlets

sedimentary structures and sand body geometry: examples from

North Carolina. South Carolina and Georgia. J. Sediment. Petrol.

49:1073-1092.
kennish, M. J. & R. K. Olsson. 1975. Effects of thermal discharges

on the microstructural growth of Mercenaria mercenaria. Environ.

Geol. 1:41-64.
Kreauter, J. N. & M. Castagna. 1980. Effects of large predators on

the field culture of the hard clam, Mercenaria mercenaria. U.S.

Natl. Mar. Fish. Serv. Fish. Bull. 78:538-541.
Mackenzie, C. L., Jr. 1977. Predation on hard clam (Mercenaria

mercenaria) populations. Trans. Am. Fish. Soc. 106:530-537.
Menzel, R. W. & H. W. Sims. 1964. Experimental farming of hard

clams, Mercenaria mercenaria, in Florida. Proc. Natl. Shellfish.

Assoc. 53:103-109.
Menzel, R. W., E. W. Cake, M. L. Haines, R. E. Martin & L. A. Oslen.

1976. Clam mariculture in northern Florida: field study on

predation. Proc. Natl. Shellfish. Assoc. 65:59-62.
Peterson, C. H. 1982. Clam predation by whelks (Busycon spp.):

Experimental tests of the importance of prey size, prey density

and seagrass cover. Mar. Biol. (Woods Hole) 66:159-170.
Rhoads, D. C. & G. Panella. 1970. The use of molluscan shell growth

patterns in ecology and paleoecology. Lethaia 3:143-161.
Walker. R. L. 1981. The distribution of the Atlantic oyster drill,

Urosalpinx cinerea (Say), in Wassaw Sound, Georgia. Ga. J. Sci.

39:126-139.
. 1982. The gastropod, Thais haemastoma, in Georgia:

T. h. floridana or T. h. canaliculata? Gulf Res. Rep. 7:183-184.
__^_ . (in preparation! Population dynamics of the hard clam.

Mercenaria mercenaria (Linne), and its relation to the Georgia
hard clam fishery. Atlanta, GA: Georgia Inst, of Technology;
Thesis.
. (in press) Effects of density and sampling time on the

growth of the hard clam, Mercenaria mercenaria, planted in
predator-free cages in coastal Georgia. Nautilus 98.
, M. A. Fleetwood & K. R. Tenore. 1980. The distribution

174 Walker

of the hard clam, Mercenaria mercenaria (Linne), and clam Whetstone, J. M. & A. G. Eversole. 1977. Predation on hard clams,

predators in Wassaw Sound, Georgia. City, State: Ga. Mar. Sci. Mercenaria mercenaria, by mud crabs, Panopeus herbstii. Proc.

Cen. Tech. Rep. 80-8;59 p. Natl. Shellfish. Assoc. 68:42-48.

Walker, R. L. & K. R. Tenore. (in press) The distribution and . 1981. Effects of size and temperature on mud crabs,

production of the hard clam, Mercenaria mercenaria. in Wassaw Panopeus herbstii, predation on hard clams, Mercenaria mercenaria.

Sound, Georgia. Estuaries 1. Estuaries 4: 153—156.

Journal of Shellfish Research, Vol. 3, No. 2, 175-182, 1983.

WATER QUALITY FLUCTUATIONS IN RESPONSE TO VARIABLE LOADING
IN A COMMERCIAL, CLOSED SHEDDING FACILITY FOR BLUE CRABS

DON P. MANTHE 1 , RONALD F. MALONE 1
AND HARRIET M. PERRY 2

1 Department of Civil Engineering
Louisiana State University
Baton Rouge, LA 70803

2 Gulf Coast Research Laboratory
Ocean Springs, Mississippi 39564

ABSTRACT A commercial, closed, recirculating seawater facility using biological filters for control of nitrogenous
metabolites is described. The volume of each system was 7,560 v. Loading densities of over 1,000 crabs (Callineetes sapidus
Rathbun) were maintained in each system. Water quality parameters (NH 3 -N, N0 2 -N, NO3-N, pH, dissolved oxygen,
salinity, temperature, alkalinity) affecting crab survival at molting were monitored for a 2-month period, and safe opera-
tional ranges were established. Alkalinity and pH values declined in the systems, demonstrating a limited buffering capacity.
Values of NO3-N exceeding 350 mg/C were observed with no apparent effects to the crabs. Increased molting mortality
was observed when concentrations of nitrite approached 1.6 mg/C N0 2 -N. Nitrite accumulations were associated with
depressed oxygen levels which were induced by peak system loadings or equipment failure. Successful molting rates in
excess of 957c were achieved at nitrite and ammonia concentrations below 1 mg/P.

KEY WORDS Aquaculture, biological filter, blue crab, Callineetes sapidus, closed system, molting, water quality

INTRODUCTION

Reported landings for soft-shell crabs have declined
drastically in most states harvesting the resource (Jaworski
1971 ; Otwell et al. 1980;Perry, Ogle and Nicholson 1982).
According to Jaworski (1971) the reduced production is
attributed to a deterioration of coastal zones and accom-
panying decline in water quality. Despite the decline in
landings, the value of soft-shell crabs has continued to rise
in the Gulf of Mexico area and averages S4.50 Kg" 1 as
compared to SI. 94 Kg" 1 in 1970 (Perry et al. 1982).

Traditionally, premolt blue crabs were collected and
held in natural waters in floating boxes or pens until they
molted (Haefner and Garten 1974). The continuing decline
in coastal water quality and subsequent increase in mortality
of molting crabs have forced fishermen to turn to onshore
facilities to reduce crab losses during molting (Jaworski
1982). The potential value of using closed, recirculating
seawater systems for maintaining molting crabs has been
demonstrated by a few successful commercial operators
(Perry, Ogle and Nicholson 1982). Recirculating systems
reduce labor requirements and eliminate the exposure of
crabs to deleterious environmental effects during the vulner-
able molting period; however, their success has been
marginal because of the lack of established design criteria
and management guidelines (Van Gorder and Fritch 1980,
Ogleetal. 1982).

In the Gulf of Mexico, where crab fishermen are often
limited by the availability of premolt blue crabs, shedding
operators strive for a molting mortality of less than 5% to
maintain production and commercial viability. On the
eastern coast of the United States, commercial shedding

systems generally have access to an abundance of crabs
and, therefore, can absorb a higher crab mortality (5 to
40%).

The operation of a successful, closed, recirculating
aquaculture system depends on the maintenance of accept-
able water quality. Wheaton (1977) and Spotte (1979)
summarized the ability of biological filters in the closed
systems to convert ammonia (NH 3 ), the principal nitro-
genous excretory metabolite of Crustacea (Hartenstein
1970), to the relatively nontoxic nitrate (N0 3 ) by bacterial
nitrification.

In 1982. a project was initiated to establish production
levels and operating parameters for closed, recirculating
seawater systems currently used to hold shedding crabs.
This approach provided a unique opportunity to comple-
ment experimental research on molting crabs (Manthe et al.
in press) with direct observations and data from the com-
mercial sector. In this report we describe the influence
of commercial operating procedures on water quality in a
large-scale shedding facility.

MATERIALS AND METHODS

Description of Commercial Facility

The commercial shedding operation was located in an
uninsulated building in LaCombe, LA. The facility con-
sisted of two separate systems (Figure 1 ), each with eight
holding tanks, two biological filters, one algal tank, and a
reservoir. The reservoir was located outside the building
and was partially buried in the ground. The facility is a
modification of one described by Perry. Ogle and Nicholson

175

176

Manthe. Malone and Perry

CRAB TANK

-Mr

BIOLOGICAL FILTER

ALGAE FILTER

RESERVOIR

CRAB TANK Vf

BIOLOGICAL FILTER

"Lj

Figure 1. Schematic diagram of one of the commercial systems.

(1982). Descriptions and dimensions of the fiberglass tanks
are presented in Table 1 .

TABLE 1.
Dimensions of Commercial System.

Water
Length Width Depth Depth Area Volume
Description (m) (m) (m) (m) (m 2 ) (m )

Crab tank

2.44

1.07

0.30

0.13

2.61

0.34

Biological

Filter

2.44

0.91

0.30

0.24

2.22

0.53

Shell

Filter Bed

2.29

0.91

0.08

—

2.08

0.17

Dolomite

Filter Bed

2.29

0.91

0.04

—

2.08

0.08

Carbon

Filter Bed

2.29

0.91

0.03

2.08

0.06

Algal Filter

2.44

0.91

0.30

0.24

2.22

0.53

Holding

Reservoir

2.74

1.37

1.52

1.07

3.75

4.02

Water levels in the crab tanks were controlled by 12.7-cm
standpipes constructed from 3.2-cm polyvinylchloride (PVC)
pipe. Water input to the discharge nozzle in the crab tanks
consisted of capped 1.3-cmPVC pipe with two 0.3 cm holes
to promote active aeration in the tanks.

The biological filters were constructed with a fiberglass
partition that was positioned 15.2 cm from one end of the
tank with holes drilled in the lower 2.5 cm of the partition.
The head chamber received the overflow from the crab tanks

through the standpipe that discharged beneath the water
level in the head chamber. The function of the head
chamber was to direct water flow under the submerged,
updraft, biological filter. The biological filter consisted of
a 7.6-cm layer of washed clam shells (Rangia cuneata)
(2- to 3-cm diameter) on the bottom, overlaid by 3.8 cm of
dolomite (3-mm grain size), and 2.5 cm of activated carbon
(1- to 3-mm grain size) (Figure 2). Each layer of medium was
separated by nylon window screen. The filter bed rested
on 1 .3-cm egg crate louvering supported by 2.5-cm PVC
pipe lengths. The water level in the biological filter was
approximately 5 cm above the top of the activated carbon
bed; overflow to the algal tank was provided through a
5.1-cm PVC pipe. Theoretically, the biological filters per-
formed two main functions: mineralization and nitrifica-
tion. These functions take toxic nitrogen waste products
produced by the crabs and convert them to relatively
nontoxic forms. By design rational, buffering was accom-
plished using carbonate filtrants (shell and dolomite), and
physical adsorption of dissolved organic carbon occurred
on activated marine carbon.

The algal tank contained 1 1 baffles that alternately
extended to within 7.6 cm of the tank sides. Attached algae
grew on the sides of the baffles and water flowed in a
serpentine fashion from one end of the tank to the other.
Another 5.1-cm PVC pipe, from the other four crab tanks
and biological filter in each system, drained into the algal
tank half way through each filter. Each algal filter was
illuminated by two 1.2-m fluorescent fixtures which
contained four 40-W Grow Lux® lights. Plants in the

Water Quality Fluctuations in a Commercial Shedding Facility

177

Figure 2. Cross section of biological filter.

systems included water milfoil (Myriophyllum sp.) that
floated on the algal filter surface, and attached filamentous
green algae on the filter walls. No algae were harvested from
either of the systems and the algal filters were provided
with a constant light regime. Water flowed by gravity from
the algal filters and was carried by 7.6-cm PVC pipe through
the wall of the building to the partially buried reservoir
outside. The algal filters were incorporated into the system
to remove nitrate, the end product of nitrification.

The large reservoir helped to buffer any rapid changes
in water quality in the systems. Rapid water quality changes
(typically transitional increases in ammonia and nitrite)
can be associated with the introduction of a large number
of crabs to a system that has been acclimated to a smaller
number of crabs, a common practice in commercial opera-
tions. Water was constantly circulated from the reservoir
via a 5.1 -cm PVC pipe to a 0.25 kW pump (model Dayton®
6K695). The pipe was screened to prevent the intake of
large debris. The water was then distributed through a
3.2-cm overhead PVC pipe to the crab tanks by a series of
1.9-cm PVC valves and tees. Water was sprayed under
pressure into the crab tanks at a constant (total system)
flow rate of 83.3 2/min.

Methods

Artificial seawater (Rila Mix®) was used in the commer-
cial facility. The two systems were constructed and operated
one year prior to the study and had been shut down in the
winter of 1982—83. Start up in March of 1983 consisted
of turning on the pumps and diluting the systems to volume
with fresh well water. Fresh water was added to the systems
to offset evaporation, but no water changes were made
during the period of observation. Intermolt blue crabs and
miscellaneous estuarine fish were used to acclimate the

biological filters until April. During the study, premolt blue
crabs ranging from 10 to 15 cm in carapace width were
taken from Lake Pontchartrain, LA, and held in the systems.
Vinyl-coated, wire-mesh enclosures 0.3 cm in diameter
isolated crabs that had molted. System management
included visual inspections every 3 to 4 hours to collect
soft-shell crabs and to remove mortalities, debris, and
exuviae. Crabs in the shedding systems were not fed at any
time. Mean residence time for a typical crab in the system
was estimated to be 7 days.

Systems were monitored at 9:00 a.m. each day for
temperature, salinity, dissolved oxygen, ammonia, nitrite,
and pH. A 1 £ sample of water was taken from each system
and analyzed immediately for total ammonia and nitrite.
Determinations of alkalinity levels and nitrate concentra-
tions were made on a weekly basis. Techniques and instru-
mentation to measure the above-mentioned parameters
are listed in Table 2. Crab densities in each system were
recorded daily after crab additions were made in the after-
noon (2:00 p.m.). Data were collected through the spring
shedding season of 1983, and systems were numbered
( 1 and 2, respectively) for reporting and identification
during the interpretation of results.

RESULTS AND DISCUSSION

pH, Alkalinity, Salinity, Nitrate

Water quality observations for pH. alkalinity, and nitrate
are illustrated in Figure 3. Both systems behaved similarly
with regard to the monitored parameters. Each system
initially exhibited a pH of 7.7, and declined to values
between 7.0 and 7.2. This reduction of pH was associated
with a decline in alkalinity. The systems displayed initial
alkalinities of 70 mg/2 CaC0 3 , declining to values as low as

178

Manthe, Malone and Perry

TABLE 2.
Measurements Taken and Techniques.

Parameter

Instrument or Test

Reference

Total ammonia

Orion 95-10 ammonia electrode/

APHA (1980)

asNH 3 -N

Orion 701 A digital ionalyzer

Nitrite as

Bausch and Lomb Spectronic 20,

Sulfanilamide-

N0 2 -N

Spectrophotometer

based colori-
metric reaction
APHA (1980)

Oxygen as

Yellow Springs Instrument Co.

Dissolved oxygen meter. Model 51

Salinity

American Optical Refractometer

Mini (Model 47) pH meter

Nitrate as

Modified hydrazine reduction

Spotte(1979)

NO3-N

Alkalinity as

Titration

APHA (1980)

CaC0 3

30 mg/C CaC0 3 , suggesting a limited capability of the
dolomite and shell layers to buffer pH changes. These
findings are consistent with the observations of Bower
et al. (1981), who noted the limited ability of calcareous
filtrants to maintain pH above 8.0. In this study, pH values
fell within the 7.0 to 8.5 range of optimum nitrification
rates for biological filters (Wheaton 1977), although filters
can be acclimated to pH values lower than 7.0 (Haug and
McCarty 1972). We conclude that the dolomite/shell bed
was sufficient for control of pH above 7.0 even after two
years of operation with no filter maintenance, and that this
pH apparently does not adversely affect the crabs. In fact,
the lower pH may be beneficial in that it reduces ammonia
toxicity because of the equilibrium reaction between NH*,
and NH 3 (Wheaton 1977, Spotte 1979).

The concentration of nitrates increased throughout the
study and was directly related to the increased crab loadings
of both systems. Nitrate levels in systems 1 and 2 were
171 and 214 mg/£ N0 3 -N, respectively, at the beginning of
the observation period. Differences in these accumulated
nitrate concentrations apparently resulted from unequal
numbers of crabs in each system and the total time that
the individual systems were operated in the year prior to
the study (system 2 was operated for a longer period
in 1982). Observations of the algal filters revealed little or
no algal growth despite efforts to reintroduce algae from
local sources and another commercial system. Values
exceeding 350 mg/C N0 3 -N were observed at the end of
this study with no apparent deleterious effects. Nitrate is
generally not toxic to marine organisms even at elevated
levels (Hirayama 1974. Siddall 1974). Salinity in the
systems remained constant at 4 and 5 ppt in systems 1 and
2, respectively.

Ammonia, Nitrite, Temperature, Dissolved Oxygen

Ammonia and nitrite increases were closely related to

increases in crab concentrations (Figure 4). and both
systems showed comparable results in terms of water
quality and crab loadings. In system 1, total ammonia
concentrations remained under 0.4 mg/C NH 3 -N regardless
of crab density. Nitrite concentrations, however, increased
to 1.6 mg/£ N0 2 -N during a period of heavy loading.
During the heaviest crab loading in system 2, ammonia
levels approached 1.0 mg/C NH 3 -N, with a similar increase
in nitrite. On May 5, increased mortality of molting crabs
was observed in system 1 when nitrite concentrations
approached 1.6 mg/2 N0 2 -N. A pump malfunction occurred
in system 2 on May 28, and nitrite concentrations subse-
quently increased. Concentrations above 1.2 mg/2N0 2 -N
were observed on May 29, but returned to low levels the
following day. Mevel and Chamroux (1981) found that
during similar pump malfunctions nitrate levels decreased
and nitrite levels increased. They concluded that nitrate
was reduced to nitrite when oxygenation of the environ-
ment was deficient, and that bacteria were responsible for
the dissimilatory nitrate reduction. This might explain the
increase of nitrite observed in our study during the pump
malfunction.

Figure 5 illustrates the temperature and dissolved
oxygen levels recorded in the systems under study. Water
temperature in the systems equilibrated rapidly to ambient
air temperatures which varied from 11° to 27°C. Higher
temperatures decreased the overall carrying capacity of the
systems. This was supported by the inverse effect of
temperature on the saturation levels of dissolved oxygen
(Figure 5). Higher temperatures also increased the metabolic
rates of both the heterotrophic and nitrifying bacteria
(Wild et al. 1971) and the crabs (Laird and Haefner 1976).
Oxygen levels in both systems were influenced by this
increased biological activity. Oxygen measurements in the
commercial systems were taken between the biological
filter and the algal tank. Because of the large surface area
on the top of the upflow biological filters, some surface
reaeration may have occurred upstream of the dissolved
oxygen measurement point; however, in both commercial
systems the lowest dissolved oxygen values were concurrent
with peak values of nitrite and crab loadings, thereby
suggesting intense activity in the biological filter. These
observations were consistent with those of Manthe et al.
(in press) which identified dissolved oxygen as the factor
limiting the efficiency of nitrification beds in experimental
crab shedding systems of the same design. That study also
demonstrated that as dissolved oxygen concentration
decreased, toxic nitrite concentrations increased and
significant crab mortality occurred.

In the latter part of this study, the biological filters
began to overflow in the head chambers. Accumulations of
detritus were observed in the nylon screens separating the
different layers of medium in the filter bed. These accumula-
tions may have led to the short circuiting of the filter bed and
thus reduced its nitrification ability. Annual breakdown

Water Quality Fluctuations in a Commercial Shedding Facility

179

10 r

9 -

£'

5 -

SYSTEM I

SYSTEM 2

I i I I I 1 1 1 1 1 1 L

3/24 28 4/1 5 9

■ i i i i — i — i — i — i — i—

3 17 21 25 29 5/1 5 9

i i i i — i — i — i — i — i

13 17 21 25 29

c 80 -

o» 70 -
E

rp 60

o 50
O

w 40

t 30

3 20

2. .o

ii i i i — i — i

i i i i i i i I I I I 1 1 L

J I

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5 9 13 17 21 25 29

360 r

g 320

2 280

240

O 200
u

160

i i i i i i i i i i — i — i — i — i — i — i — i i

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5 9 13 17 21 25 29

DAYS

Figure 3. Supporting water quality parameters for the commercial systems (pH, alkalinity, and nitrate).

180

Manthe. Malone and Perry

12:50

1000

750

500

250

System I

I I I 1 1 I 1 L-

_l I I l_

_l 1 I I I

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5
System I

13 17 21 25 29

AMMONIA
NITRITE

3/24 28 4/1 5 9
1250 |~ System 2

1000

i i i i

-2.0

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5
System 2

9 1 .6

z I 2

cc 08

<-> 04

13 17 21 25 29

AMMONIA
NITRITE

13 17 21 25 29

00
3/24 28 4/1 5 9 13 17 21 25 29 5/1 5

DAYS

Figure 4. Ammonia and nitrite concentrations in relation to crab densities (systems 1 and 2).

Water Quality Fluctuations in a Commercial Shedding Facility

181

— 15

I-
<

System

TEMPERATURE
2

I I I I I I I I I I I I I I 1 I I I I I I I 1 1 1 1 1 1 1 1 1 1 L.

-|I2
10
8
6
4
2

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5 9 13 17 21 25 29 O

System 2

10 I I I I I I I L

I ' I I I 1 1 1 I 1 1 1 1 1 L.

I ' ' I L

10
8
6
4
2

3/24 28 4/1 5 9 13 17 21 25 29 5/1 5 9 13 17 21 25 29

DAYS

Figure 5. Temperature and dissolved oxygen values (systems 1 and 2).

UJ
O

z
o
o

and cleaning of the biological filter should be considered
when using this design.

Throughout the study, acceptable water quality was
maintained with this filter design and successful molting
rates of more than 95% were observed in the facility.
Loading values of over 1 ,000 crabs were maintained by each
system over the observed period. Table 3 summarizes
observed operational ranges for selected water quality
parameters in the systems studied.

TABLE 3.

Observed operational ranges for selected
water quality parameters.

Parameter

Range

Total ammonia

Nitrite

Nitrate

Temperature

- 1 mg
0- 1 mg
- 350 mg
7-8
11°-27°C

sr 1

Decreased molting success was observed when concen-
trations of nitrite approached 1.6 mg • T 1 N0 2 -N. Total
ammonia levels did not rise above 1.0 mg • C" 1 NH 3 -N and
these levels had no apparent harmful impact on the molting
crabs. In both systems the lowest dissolved oxygen values
were concurrent with peak values of crab density and nitrite,
indicating an intense oxygen demand in the biological
filters to process the increased production of nitrogenous
waste. Monitoring of nitrite and dissolved oxygen concentra-
tions appear to be of critical importance to commercial
softshell production in closed systems.

ACKNOWLEDGM ENTS

This research was supported by the Louisiana Sea Grant
College Program. Collaborative support was provided by the
Mississippi-Alabama Sea Grant Consortium. These programs
are elements of the National Sea Grant Program, under the
direction of NOAA, U.S. Department of Commerce. We
gratefully acknowledge Drs. Ronald Becker and James I. Jones

182

Manthe, Malone and Perry

for their encouragement and supportive services; and Dr.
Edwin W. Cake, Jr., for his editorial reviews. The authors
also express their thanks and gratitude to Mr. Cultus
Pearson of LaCombe, LA, for sharing his knowledge of crab
shedding systems and for permitting access to his systems

for our study. Housing accommodations and research
facilities during the study were provided by the Louisiana
Department of Wildlife and Fisheries. References to trade
names do not imply product endorsement by the authors
or by the National Sea Grant Program.

REFERENCES CITED

American Public Health Association (APHA). 1980. Standard
Methods for the Examination of Water and Wastewater. 15 ed.
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Pollution Control Federation, American Public Health Associa-
tion. 1,134 p.

Bower, C. E., D. T. Turner & S. Spotte. 1981. pH maintenance in
closed seawater systems: limitations of calcareous filtrants.
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Haefnei, P. O., Jr. & D. Garten. 1974. Methods of handling and
shedding blue crabs, Callinectes sapidus. Va. Inst. Mar. Sci.
Mar. Res. Adv. Ser. 8:1-14.

Hartenstein, R. 1970. Nitrogen metabolism in non-insect arthropods.
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Haug, R. T. & P. I. McCarty. 1972. Nitrification with submerged
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Hirayama, K. 1974. Water control by filtration in closed systems.
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. 1982. History and status of Louisiana's soft-shell blue

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Laird, C. E. & P. A. Haefner, Jr. 1976. Effects of intrinsic and
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Manthe, D. P., R. F. Malone & S. Kumar, (in press) Limiting factors
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Aquacult. Engineer.

Mevel. G. & S. Chamroux. 1981. A study on nitrification in the

presence of prawns {Penaeus japonicus) in marine closed systems.

Aquaculture 23:29-43.
Ogle. J. T., H. M. Perry & L. C. Nicholson. 1982. Closed recirculating

seawater systems for holding intermolt crabs: literature review,

systems design and construction. Ocean Springs, MS: Gulf Coast

Research Laboratory Tech. Rep. Ser. 3:1—11.
Otwell, W. S., J. C. Cato & J. G. Halusky. 1980. Development of a

soft crab fishery in Florida. Fla. Sea Grant Coll. Rep. 31:1-56.
Perry, H. M., J.T. Ogle & L. C. Nicholson. 1982. The fishery for soft

crabs with emphasis on the development of a closed recirculating

seawater system for shedding crabs. Perry, H.M.andW.A.Van Engel,

eds. Proceedings of the Blue Crab Colloquium; 1979 October

16-19; Biloxi, MS: Gulf States Mar. Fish. Comm. Ann. Meet. 7:

137-152. Available from GSMFC, Ocean Springs, MS.
Perry, H. M., G. Adkins, R. Condrey, P. Hammerschmidt, S. Heath,

J. Herring, C. Moss, G. Perkins & P. Steele. 1982. A profile of

the blue crab fishery of the Gulf of Mexico. Ocean Springs, MS:

Gulf States Marine Fisheries Commission. Compl. Rep., Contr.

No. 000-010: 184 p.
Siddall, S. E. 1974. Studies of closed marine culture systems.

Prog. Fish-Cult. 36:8-15.
Spotte, S. 1979. Fish and Invertebrate Culture, Water Management

in Closed Systems. New York, NY: John Wiley & Sons, Inc.

179 p.
Van Gorder, S. D. & J. D. Fritch. 1980. Filtration techniques for

small-scale aquaculture in a closed system. Proc. Annu. Conf.

Southeast. Assoc. Fish Wildl. Agencies 34:59-66.
Wheaton, F. W. 1977. Aquacultural Engineering. New York, NY:

John Wiley & Sons, Inc. 708 p.
Wild, H. E., Jr., C. N. Sawyer & T. C. McMahon. 1971. Factors

affecting nitrification. J. Water Pollut. Control Fed.

43:1845-1854.

Journal of Shellfish Research, Vol. 3, No. 2, 183-193, 1983.

BLUE CRAB (CALLINECTES SAPIDUS RATHBUN) POPULATIONS IN MID-CHESAPEAKE BAY
IN THE VICINITY OF THE CALVERT CLIFFS NUCLEAR POWER PLANT, 1968-1981

GEORGE R. ABBE

Academy of Natural Sciences

Benedict Estuarine Research Laboratory

Benedict, Maryland 20612

ABSTRACT Population data of the blue crab Callinectes sapidus Rathbun were collected from 1968 to 1981 to deter-
mine the affects of waste heat from the Calvert Cliffs Nuclear Power Plant on abundance, size distribution, sex ratio, and
seasonality of occurrence. Crabs were sampled using commercial crab pots of 25-mm-mesh set within (Plant Site) and out-
side (Kenwood Beach and Rocky Point) the main area of thermal influence. Five pots per station were fished four days
per week during alternate weeks from May to November or December. Crabs were sexed, measured, and weighed by sex.
In 14 years, a total of 10,600 pot samples yielded 57,078 crabs (5.38 per pot) of which 74.1% were legal size (^ 127 mm
carapace width) and 5 1.5% were males. During seven pre-operational years (1968-74), the number of crabs per pot averaged
4.15 at Kenwood Beach (32.6%), 4.12 at Plant Site (32.4%), and 4.46 at Rocky Point (35.0%). During seven operational
years (1975-81), the number of crabs per pot averaged 6.24 at Kenwood Beach (33.3%), 6.37 at Plant Site (34.0%), and
6.15 at Rocky Point (32.8%). Increased catch during the operational period was due to extreme abundance in 1981 when
the mean catch was nearly 17 crabs per pot. Ten population variables were tested for differences between pre-operational
and operational periods and among stations and years. Data analyses revealed many differences among years due to natural
fluctuations in the size and structure of Chesapeake Bay blue crab populations. Two station differences were detected;
males at Kenwood Beach were slightly larger than at the other stations (p <0.01), and percent males at Kenwood Beach
was higher than at Rocky Point (p < 0.01). The overall similarity of stations and periods indicates no evidence of any
detrimental effect on the crab populations caused by operation of the Calvert Cliffs Nuclear Power Plant.

KEY WORDS: blue crab, Callinectes sapidus, Calvert Cliffs, Chesapeake Bay, nuclear power plant, thermal effluent

INTRODUCTION

For nearly a century the blue crab Callinectes sapidus
Rathbun has been the basis of one of the major commercial
fisheries in the Chesapeake Bay area. From the late 1940's
to the late 1950's the annual catch averaged nearly 27.2 X
10 6 kg(60X 10 6 lb) valued at S3 million (Van Engel 1958).
From 1968 to 1975 annual landings increased to almost
29.9 X 10 6 kg (66 X 10 6 lb) valued at $7.7 million (U.S.
Fish and Wildlife Service 1970a, b; National Marine Fish-
eries Service 1972- 1976a, b). From 1976 to 1980 mean
landings fell to 26.3 X 10 6 kg (58 X 10 6 lb), but dockside
value increased to $13 million (National Marine Fisheries
Service 1977-1979a, b, 1980, 1981, 1982). Record landings
occurred in 1981 with 46.3 X 10 6 kg (102 X 10 6 lb) valued
at $27 million (National Marine Fisheries Service 1982).

The size and economic importance of this fishery are
obvious cause for concern regarding industrial construction
which could affect it. Mihursky and Kennedy (1967) dis-
cussed problems associated with heated discharges from
power plants including the fact that many plants discharge
water heated to 38-46°C. Tagatz (1969) also indicated
that power plant discharges of heated waste water might
be a threat to blue crabs.

In 1968 Baltimore Gas and Electric Company began
construction of the Calvert Cliffs Nuclear Power Plant
(CCNPP), a two-unit generating station located on Chesa-
peake Bay in Calvert County, Maryland, about 15 km
north of the mouth of Patuxent River. Bay water is used in
a once-through cooling system at a rate of 9.08 X 10 6 £/min,
heated 6.7°C (maximum) above ambient and discharged at
2.7 m/sec through a high-velocity jet 260 m from shore
(Baltimore Gas and Electric Company 1970). Mixing of the

discharge and receiving water is rapid and the area enclosed
by the +2°C isotherm is only 0.34 km 2 assuming 10%
recirculation (Academy of Natural Sciences of Philadelphia
ct al. 1980); however, thermal increases of 0.5 to 1.0°C
above ambient have been detected more than 3 km away.

In addition to being one of the most abundant commer-
cial species in the Chesapeake Bay, the blue crab is also one
of the most tolerant of a wide range of salinities and
temperatures. Tagatz (1969) has shown that at salinities of
7 ppt, somewhat lower than the 10 to 15 ppt at Calvert
Cliffs, 50% of the crabs acclimated to 22°C will survive
48 hours at a temperature of 36.9°C. Burton (1978)
exposed juvenile blue crabs acclimated at 15 and 25°C to a
rapid 10°C increase, held them at the elevated temperature
for four minutes, and returned them to the acclimation
temperature over a 15-minute decay period. Weight-specific
oxygen consumption indicated that responses were due to
normal physiological temperature compensation and not to
thermal stress. He concluded that increases of up to 10 C
would have minimal adverse seasonal effect on blue crabs
when exposure time was held to 20 minutes. Because
maximum temperatures near the CCNPP discharge are
several degrees below 37°C and because blue crabs are
strong swimmers capable of relatively rapid movement,
mortalities resulting from the thermal discharge were not
expected. Sublethal temperatures, however, could affect
the distribution or structure of the total population, so
that numbers of crabs, their mean sizes or sex ratios would
be abnormally altered. Because large fluctuations in annual
abundance of blue crabs have been well documented

183

184

ABBE

(Pearson 1948, Van Engel 1958, Tagatz 1965), this study
was designed to examine abundance, seasonality of occur-
rence, sex ratio, and size-frequency distribution of the crab
populations in the vicinity of the CCNPP over several
years, and to ascertain whether any significant changes in
these parameters have resulted from plant operation. The
plant became operational in early 1975 and Unit 1 began
commercial production in May 1975; Unit 2 began opera-
tion in 1977. Thus seven years of pre-operational data and
seven years of operational data were collected from 1968 to
1981. This paper reports the effects of power plant opera-
tion on the local crab populations and provides descriptive
statistics of these populations over a 14-year period.

MATERIALS AND METHODS

Stations

The center of the study area was adjacent to the Calvert
Cliffs plant site located approximately 7.6 km northwest of
Cove Point on the western shore of Chesapeake Bay (Figure 1).
Although this station was within 100 m of the discharge, it
did not receive the full impact of the thermal plume.
Temperatures averaged 1° to 2°C above ambient during
operational years; water depth was approximately 2.5 m.
The upper station was located near Kenwood Beach, 6.4 km
from the discharge at 3.7 to 4.6 m water depth; the lower
station was southeast of Rocky Point 3.8 km from the
discharge at 3.5 m water depth. The Kenwood Beach and
Rocky Point stations were outside of the predicted area of
thermal affect when they were established in 1968. Plant
operation, however, did result in occasional temperature
increases of up to 1°C at Rocky Point; Kenwood Beach
remained unaffected. Salinity at the Rocky Point station
averaged 1 to 2 ppt higher than at Kenwood Beach.

Figure 1. Locations of crab pots in mid-Chesapeake Bay from 1968
to 1981.

Study Design

Commercial potting techniques (Van Engel 1962) and
crab pots of 25-mm-mesh were used to sample the crab
populations at the three stations from spring (generally
early May) until late fall when water temperatures declined
to levels at which crabs became inactive (10-12°C in
November or December). Commercial crab pots are generally
constructed of 38-mm-mesh and will hold few crabs less
than 76 mm (3 in.) in carapace width; however, the smaller
mesh pots used in this study allowed some crabs less than
51 mm (2 in.) to be caught.

During alternate weeks throughout the season, five pots
(three in 1968) at each station were baited daily with
menhaden and fished for four successive days. Station
catches were weighed by sex and all crabs were measured
to the nearest one-eighth-inch (3 mm). Field measurements
were later converted to metric.

Bottom temperature and salinity were determined
monthly by thermistor probe and titration, respectively,
from 1968 to 1978 and daily during the weeks fished from
1979 to 1981 with a Beckman RS5-3 portable salinometer.
Dissolved-oxygen concentrations were determined monthly
through 1974 and daily thereafter either by Winkler titra-
tion or with a YSI Model 57 dissolved-oxygen meter.

Statistical Analysis

A cross-nested analysis of variance (Hicks 1973) was
used to compare various parameters of the crab populations
and thus test for differences between the pre-operational
and operational periods. Population parameters included
the number caught per pot for legal size crabs and for total
crabs, mean widths and weights of males, females, and
combined sexes, the percent legal size crabs, and percent
males. The crossed effects in this analysis were station and
year and the nested effect was year within period. The
period effect was tested against the year (period) error term
while other effects were tested against the highest order
interaction term for this model. All parameters except
percent legal size and percent male were yearly averages;
because averages of large samples tend to be normally
distributed, no transformation was required. The percent
legal size and percent males at each station for each year
were transformed by arcsine-squareroot transformation to
stabilize variances and improve normality (Thoni 1967).

Percent males were also examined over the entire 14-year
period by analysis of covariance (Hicks 1973) using logit-
transformed data (Cox 1970).

RESULTS AND DISCUSSION

Summaries of the annual blue crab catches made in the
Calvert Cliffs area during 1968—1974 (pre-operational) and
during 1975—81 (operational) are presented in Tables 1 and
2, respectively. In 14 years of study. 10,600 pot samples
produced 57,078 crabs (5.38 per pot) of which 51 .5% were
males and 74.1% were legal size (^ 127 mm carapace width).

Blue Crab Populations in Mid-Chesapeake Bay

185

TABLE 1.

Summary of abundance, size, and sex composition of blue crab catches near the Calvert Cliffs
Nuclear Power Plant from 1968 to 1974 (preoperational period).

1968

1969

1970

1971

1972

1973

1974

Grand Mean

Total number

239

2,833

1,493

4,792

3,041

3,059

3,970

2,775

Number of males

158

1,995

914

2,65 7

1,794

1,753

2,366

1,662

Number of females

838

5 79

2,135

1,247

1,306

1,604

1,113

Percent males

66.1

70.4

61.2

55.4

59.0

57.3

59.6

61.3*

Total weight (kg)

367

228

709

448

479

630

416

Mean weight per crab (g)

201

130

153

148

147

157

159

155*

Male weight (kg)

262

145

417

277

295

400

261

Mean weight per male (g)

209

131

159

157

154

168

169

164*

Female weight (kg)

106

293

171

185

230

155

Mean weight per female (g)

185

126

143

137

142

143

145*

Number of legal size crabs ( > 127 mm)

206

2,006

1,128

3,629

2,195

2,388

2,942

2,071

Number of sublegal size crabs

827

365

1,163

846

671

1,028

705

Percent legal

86.2

70.8

76.5

75.7

72.2

78.1

74.1

76.2*

Mean crab width (mm)

153

134

140

138

137

143

141

141*

Mean width of males (mm)

151

134

139

135

134

141

139

139*

Mean width of females (mm)

157

134

142

141

146

144

144*

Total pots fished

281

472

564

760

795

898

809

654

Number of crabs per pot

0.85

6.00

2.65

6.31

3.83

3.41

4.91

3.99*

Number of legal size crabs per pot

0.73

4.25

2.00

4.78

2.76

2.66

3.64

2.97*

*Grand means of means and percents are unweighted.

TABLE 2.

Summary of abundance, size, and sex composition of blue crab catches near the Calvert Cliffs
Nuclear Power Plant from 1975 to 1981 (operational period).

1975

1976

1977

1978

1979

1980

1981

Grand Mean

Total number

Number of males

Number of females

Percent males

Total weight (kg)

Mean weight per crab (g)

Male weight (kg)

Mean weight per male (g)

Female weight (kg)

Mean weight per female (g)

Number of legal size crabs (_> 127 mm)

Number of sublegal size crabs

Percent legal

Mean crab width (mm)

Mean width of males (mm)

Mean width of females (mm)

Total pots fished

Number of crabs per pot

Number of legal size crabs per pot

4,902

2.845

2,089

3,476

5,740

3,493

15,106

5,379

2,381

1,245

1,080

1,707

3,034

1,464

6,853

2.538

2,521

1,600

1,009

1,769

2,706

2,029

8,253

2,841

48.6

43.8

51.7

49.1

52.8

41.9

45.4

47.6*

748

392

383

552

864

638

1,972

793

153

138

183

159

151

183

131

157*

384

172

217

285

478

281

863

383

161

138

201

167

158

192

126

163*

364

220

165

267

386

357

1,110

410

144

138

164

151

143

176

134

150*

4,006

1,922

1,737

2,598

4.449

2,877

10,211

3,971

896

923

352

878

1,291

616

4,895

1,407

81.7

67.6

83.1

74.7

77.5

82.4

67.6

76.4*

144

137

149

143

142

149

135

143*

140

131

148

138

143

126

138*

148

143

151

148

146

153

142

147*

902

841

756

886

879

861

896

860

5.43

3.38

2.76

3.92

6.53

4.06

16.86

6.13*

4.44

2.29

2.30

2.93

5.06

3.34

11.40

4.54*

unweighted.

*Grand means of means and percents are

186

ABBE

Considerable variation in annual population size, individual
mean size, and sex ratio is evident in data from Tables 1
and 2 with significant differences among years for all
variables examined by analysis of variance (ANOVA) (all
p < 0.01 ). There were, however, many similarities between
the two periods. For example, the annual mean number of
crabs caught per pot during the pre-operational period
ranged from 0.85 to 6.31 (a 7.4:1 ratio) and during the
operational period ranged from 2.76 to 16.86 (a 6.1 : 1 ratio).
Mean crab weights were similar during the two periods
(155 and 157 g, respectively), as were the percentages of
legal-size crabs caught (76.2% and 76.4%, respectively).
Mean carapace widths were also nearly the same at 141 mm
and 143 mm, respectively. Thus it appears that year-to-year
fluctuations were due to natural changes in population
structure and not to operation of the CCNPP.

Statistical differences among stations were detected for
only two of the ten variables tested. Male crabs were
significantly larger (p < 0.01 ) at Kenwood Beach ( 138.6 mm
carapace width) than at the Plant Site (135.6 mm) or
at Rocky Point (136.3 mm). Although these sizes differ
statistically, there is little biological significance to the
differences.

Percent males were also greater at Kenwood Beach (55%)
than at Rocky Point (48%) (p < 0.01); the 51% males at
Plant Site differed from neither. This difference probably
resulted from the higher salinities at Rocky Point than at
Kenwood Beach because the ratio of males to females
normally decreases as salinity increases (Lippson 1973).

Commercial landings in Maryland during 1968—1981
ranged from about 4.5 to nearly 27.2 X 10 6 kg (10 to 60 X
10 6 lb), while the numbers of crabs caught per pot in the
study area ranged from less than 1 to nearly 17 (Figure 2).
Linear regression analysis revealed a high correlation (r 2 =
0.88) between these two data sets indicating that crab abun-
dances near Calvert Cliffs were representative of commercial
catches in the Maryland portion of Chesapeake Bay. The
number of legal-size crabs caught per pot ranged from 0.73
to 1 1 .40 (Tables 1 and 2) and also correlated well with
Maryland landings (r 2 = 0.87).

Percents of annual legal-size crabs caught were much
more stable than abundances, ranging from 68% in 1976
and 1981 to 86% in 1968 (Tables 1 and 2). During a given
year, however, the percent of legal -size crabs caught varied
considerably. Figure 3 shows the percentage of legal-size
crabs caught by station for the weeks fished during 1981
and illustrates this seasonal variation. In May 1981 the
population consisted of crabs hatched in 1979 and 1980.
About 60% were 1979 crabs which were already of legal
size; the remainder were sublegal size from the 1980 spring
hatch. As the 1979 crabs were removed from the popula-
tion and more small 1980 crabs were recruited to the area,
the percent of legal-size crabs decreased to below 20% in
June. As the 1980 crabs grew during the season, the percent
of legal-size crabs gradually increased until a peak above

90% was reached in the fall. During November the percent
of legal-size crabs decreased again, possibly from the off-
shore movement of large crabs and from the recruitment of
small crabs hatched in early 1981 which were becoming
large enough (65 mm; Van Engel 1958) to be caught in
pots. The high correlation between the annual percent of
legal size and mean crab weight (r 2 = 0.78) is shown in
Figure 4. The lowest annual mean weights were 130 g and
131 gin 1969 and 1981, respectively, both following years
of high reproductive success. The large proportion of light-
weight, sublegal-size crabs resulted in low mean weights and
low legal-size percentages (70.8 and 67.6%). In contrast,
1968 followed a year of poor recruitment and juveniles
were scare; the mean weight was 201 g and the percent of
legal size was above 86%.

100

80 -

° 50

2
18
16
14
12
10
8
6
4
2

/'"~-~. MARYLAND and /

/ \ VIRGINIA LANDINGS /
\ A •-_ /

/
' /

V \ . / ,

MARYLAND LANDINGS / '

- f

'VA V 7 :

- /a

,\ CALVERT CLIFFS ,/\ /

/ ' '' \-- .-'"'

1 1

1 1 1 1 1 1 1 1 1 1 1

68 69 70 71 72 73 74 75 76 77 78 79 80 81

Figure 2. Commercial blue crab landings and catch per pot
Calvert Cliffs study area from 1968 to 1981.

in the

During the 14-year study period, females averaging
145 mm in carapace width were 7 mm larger than males
(138 mm). The mean weight of the males (164 g), however,
was 1 7 g more than females (147 g). This is consistent with
other studies of Chesapeake Bay (Newcombe et al. 1949),
Florida (Tagatz 1965), and Texas (Pullen and Trent 1970).
which showed that males of a given width are heavier than
females of the same width. Annual mean widths ranged
from 134 to 153 mm (Tables 1 and 2). much smaller than
the 155-, 158- and 166-mm means for crabs caught by pots
in three areas of the St. Johns River. Florida (Tagatz 1965).

Blue Crab Populations in Mid-Chesapeake Bay

187

• KENWOOD BEACH

• PLANT SITE

• ROCKY POINT

MAY JUN JUL AUG SEP OCT NOV DEC

Figure 3. Percent of catch consisting of legal-size crabs (^127 mm carapace width) at three stations during 1981.

85 -

LU
N

i 80

_l
<

75 -

UJ 70
o

°- 65

•

y=37.34+0.25x

■

•
1

i i

r 2 = 0.78

■ i i

130 140 150 160 170 180 190 200 210
MEAN CRAB WEIGHT (g)

Figure 4. Linear regression of annual percent of legal-size crabs caught and mean crab weight.

188

ABBE

Crabs in southern states apparently grow to larger sizes than
those in Chesapeake Bay. The largest crab used by
Newcombe et al. (1949) in their formulation of width-weight
curves for Chesapeake Bay crabs was 201 mm and the largest
crab from the present study was 213 mm. Of the 57,078
crabs caught near Calvert Cliffs, only 6 exceeded 203 mm
(8 in.). In contrast, Tagatz (1965) reported a 246-mm crab
from Florida and 240-mm crabs are known from Texas.

Table 3 lists numbers of males and females, their weights,
and the mean number caught per pot at each station.
Although station differences were apparent within years,
trends were similar as were overall means. Except for 1968
and 1981, the poorest and best years of the study, respec-
tively, the catch ranged from about two to seven crabs per
pot. For the 14-year study period, Kenwood Beach pots
produced a mean of 5.32 crabs per pot (32.9% of the total),
while Plant Site pots produced 5.40 crabs per pot (33.4%),
and Rocky Point produced 5.43 crabs per pot (33.6%).
These percentages are nearly identical and no statistically
significant difference exists among them (p = 0.99).

The percent of the total annual catch made at each
station ranged from 26 to 38% at Kenwood Beach, from
23 to 39% at the Plant Site, and from 30 to 45% at Rocky
Point (Figure 5). The mean number of crabs caught per pot
by station has shown no meaningful change between pre-
operational and operational periods. The overall weighted
pre-operational mean for all stations combined was 4.24
crabs per pot, whereas the weighted operational mean was
6.25 crabs per pot (4.35 if 1981 data are excluded). If 1968
data are also excluded, the pre-operational average becomes
4.52 crabs per pot. Thus, the elimination of the most- and
least-productive years yields similar long-term mean catches.
During the pre-operational period Kenwood Beach averaged
4.15 crabs per pot (32.6%), the Plant Site averaged 4.12

crabs per pot (32.4%), and Rocky Point averaged 4.46
crabs per pot (35.0%). Since 1975, these same stations
produced 6.24 (33.3%), 6.37 (34.0%), and 6.15 (32.8%)
crabs per pot. respectively; the percentages were essentially
unchanged from the pre-operational period.

Figure 6a illustrates the 14-year mean seasonality of the
crab populations by station and the similarity between these
three stations. Catches were generally small in May, as a
result of cool water temperatures (14 to 18°C) which pre-
vented full activity of the crabs. With rising water tempera-
tures, catches increased steadily until August when they
approached seven crabs per pot. A decline at all stations
was observed for September followed by a second peak in
October. The September decline resulted from a sharp
decrease in the number of males only (Figure 6b), while
females continued to increase in numbers until the October
peak was reached. Decreasing water temperatures during
November and December reduced crab activity and brought
about a rapid decline in catch size.

The October peak reflected migrating females as they
moved through the area on their way to higher salinity
water of the lower bay for eventual spawning. Females
normally spawn at salinities of 22 to 28 ppt (Sandoz and
Rogers 1944, Newcombe 1945) which larvae need to survive.
Costlow and Bookhout ( 1959) observed normal hatching at
salinities as low as 20.1 ppt with all larvae hatching as
first-stage zoeae and no prezoeae were observed. At lower
salinities, however, larvae that hatch do so as prezoeae and
do not survive (Sandoz and Rogers 1944). Although
spawning is uncommon in the Calvert Cliffs area and
although Truitt (1939) stated that sponge crabs (female
with egg pad) are seldom seen north of the Rappahannock
River in Virginia (about 80 km down-bay from the CCNPP)

LU
O

LU
Q.

^^^^^m iTMinnn nrrtni

.. i DAri/V DniklT

■

•

; ;

';[

]

68 69 70 71 72 73 74 75 76 77 78 79 80 81

Figure 5. Percent of annual catch made at each station from 1968 to 1981 based on catch per pot.

Blue Crab Populations in Mid-Chesapeake Bay

189

TABLE 3.

Numbers and weights (kg) of caught blue crabs, number of pots fished, and number of crabs caught per pot
at three stations in mid-Chesapeake Bay from 1968 to 1981.

Males

Females

Year

Number

Weight

Number

Weight

Number of Pots Fished

Number of Crabs per Pot

1968
1969
1970
1971
1972
1973
1974
1975
1976
1977
1978
1979
1980
1981
Totals

57
677
381
905
537
573
996
834
406
388
491
95 7
527
2,618
10,347

143

113

184

130

156

106

339

1,639

296

205

673

289

200

562

769

476

312

461

1.054

487

2,562

8,370

KENWOOD BEACH

99
156
192
246
265
303
276
303
275
249
284
291
283
293

1.174

3,515

0.82
6.24
3.05
6.41
3.12
2.55
5.64
5.29
3.21
2.81
3.35
6.91
3.58
17.68
5.32

1968
1969
1970
1971
1972
1973
1974
1975
1976
1977
1978
1979
1980
1981
Totals

720

212

777

602

644

743

827

409

348

630

1,002

475

2,495

9.923

117

104

121

138

102

150

311

1,489

18
270
183
630
474
580
468
708
482
360
757
776
830
2.813
9,349

PLANT SITE

63
103

60
111
108
145
373

96
156
180
257
265
325
264
300
283
254
305
295
289
300

1.314

3,569

0.59
6.35
2.19
5.47
4.06
3.77
4.59
5.12
3.15
2.79
4.55
6.03
4.52
17.69
5.40

1968
1969
1970
1971
1972
1973
1974
1975
1976
1977
1978
1979
1980
1981
Totals

598

321

975

655

536

627

720

430

344

586

1,075

462

1.740

9,131

14
78
52

157
99
77
95

116
59
64

100

172
85

212
1,381

272

191

832

484

526

574

1,044

642

337

551

876

712

2,878

9,958

ROCKY POINT

30
122

81
149

83
129
131
396

1,461

86
160
192
257
265
270
269
299
283
253
297
293
289
303
3,516

1.17
5.44
2.67
7.03
4.30
3.93
4.46
5.90
3.79
2.69
3.83
6.66
4.06
15.24

5.43

1968
1969
1970
1971
1972
1973
1974
1975
1976
1977
1978
1979
1980
1981
Totals

158
1,995

914
2.657
1,794
1,753
2,366
2,381
1.245
1,080
1,707
3,034
1,464
6,853
29,401

4,509

ALL STATIONS COMBINED

81 15 281

838 106 472

579 83 564

2,135 293 760

1,247 171 795

1.306 185 898

1,604 230 809

2,521 364 902

1,600 220 841

1.009 165 756

1.769 267 886

2,706 386 879

2,029 357 861

8,253 1.110 896

27,677 3,949 10,600

0.85
6.00
2.65
6.31
3.83
3.41
4.91
5.43
3.38
2.76
3.92
6.53
4.06
16.86
5.38

190

Abbe

L±J
CL

DQ
<

/
/

■I

— KENWOOD BEACH
— • PLANT SITE
ROCKY POINT

MAY JUN JUL AUG SEP OCT NOV DEC

Figure 6a. Monthly catch per pot by station showing the similiary between stations.

I -

MAY JUN JUL AUG SEP OCT NOV DEC

Figure 6b. Monthly catch per pot by sex showing the difference in peak abundance for males
and females.

Blue Crab Populations in Mid-Chesapeake Bay

191

except during dry years, some are occasionally seen. From
1968 to 1979. 59 sponge crabs were caught (0.34% of all
females). During the dry years of 1980 and 1981, however,
64 were collected (0.62% of all females). Salinity (20 to
21 ppt) and temperature (24 to 25°C) combinations in
late August and early September 1981 were high enough in
the study area for successful hatching although no evidence
of this was observed. Hatching this far north in the bay
would have been extremely unusual because even a blue
crab megalops in this area of the Chesapeake is rare (Cargo
1960).

Annual percentages of males are plotted by station in
Figure 7a. When stations are averaged within years, the
annual percentages show a decline from 66% in 1968 to
45% in 1981. The highest percent males occurred in 1969
(70%) and the lowest occurred in 1980 (42%). Analysis of
covariance revealed a significant decrease in percent males
since 1968 (p < 0.001). The analysis detected no difference
in the rate of decrease between stations (p >0.99); thus it
has occurred equally at all stations (Figure 7b). This decrease
in percent males might be easily explained had a long-term
increase in salinity been evident during this time, but
Figure 8 shows no increase in salinity; the regression line
is not significantly different from a no-slope line (p = 0.56).
Thus, the reasons for the decrease in male/female ratios are
not understood, nor are the implications of this decline to
a fishery in which females are worth considerably less than
males. Because choice male crabs are destined for crab
houses and restaurants to be eaten steamed, while females
and small or light males go primarily to processing plants to
be picked, the choice males may be worth two to three
times more than females during much of the season. If this
decline in percent males is more widespread than the
Calvert Cliffs area, such a decline could result in economic
losses to crabbers and others dependent on the fishery.

IOO

</)

III

r*0

KB^fl + e" 1577 " 0072 *' 1 ""]" 1

pC = r| + e -(565-0O736( f f)ll" 1
RP s [| + g" (5 «7 -0.07241 jflll" 1

68 69 70 71 72 73 74 75 76 77 78 79 80 81

Figure 7b. Curves resulting from analysis of covariance model
fitted to logit-transformed proportions of males at three stations
showing the decline in percent males and the similarity in rates
of decline.

d.0
20

J 15

SALINITY
5

i i i

y = 4.60 + 0.llx

r ! =0.03

i i i i > i i i i

69 70 71 72 73 74 75 76 77 78 79 80 81

100

o
cr
w 20

-K /'\

■ ' "'^-,/\

KENWOOD BEACH

PLANT SITE

• ROCKY POINT
. i i i i i i — i —

1 1 1 1 1 1

68 69 70 71 72 73 74 75 76 77 78 79 80 81

Figure 7a. Annual percent of catch consisting of males at the three
stations from 1968 to 1981.

Figure 8. Annual mean salinity in the Calvert Cliffs area from 1968
to 1981 showing the absence of any long-term trend. Vertical bars
represent annual salinity ranges.

Poor catches and/or dead crabs in pots were occasionally
observed during July and August as a result of low-dissolved
oxygen concentrations. Although uncommon at all stations,
these episodes occurred more often at Kenwood Beach
than elsewhere because of bathymetric differences. The
bottom at Kenwood Beach sloped from a 3- to 10-m depth
more gradually than at the Plant Site or Rocky Point
allowing anoxic water to upwell after westerly winds moved
surface waters offshore. These incidents usually lasted from
1 to 3 days when oxygen concentrations ranged from just
under 3.0 to 0.1 mg/2. Fish trapped in pots generally were
dead and crabs were dead or nearly so. Although catches at
Kenwood Beach were much reduced during these times, the
overall reduction for the season compared to other stations

192

ABBE

was minimal. May (1973) described similar occurrences in
Mobile Bay, Alabama, and discussed the responsible condi-
tions. He stated that one of the best indexes of the extent
of oxygen depletion was the mortality of fish and crabs
caught in pots.

Abundance, size, and sex ratio data indicated no special
attraction of crabs to the Plant Site station. Crabs were
attracted by warm water at the P. H. Robinson Generating
Station in Galveston Bay, Texas, during the cooler seasons
and by the entrainment of small fish (Callaway and Strawn
1975).

An estimated 4.76 X 10 6 crabs were impinged on the
rotating screens at Calvert Cliffs from 1975 to 1981. The
estimate of 3.8 X 10 s in 1980 (Hirshfield et al. 1981) was
similar to the 1975-78 mean of 4.0 X 10 s ; however, it was
well below the 1.12 X 10 6 and 1.66 X 10 6 for 1979 and
1981, respectively (Hirshfield et al. 1980, Hirshfield and
Hixson 1982). Annual impingement estimates were corre-
lated with the annual mean number of crabs caught per pot
from all stations combined (r = 0.83). Although the
number of impinged crabs was large (6.8 X 10 5 annual
mean for 1975-81), it was much lower than the estimate
of 1.95 X 10 6 crabs per year for 1976-77 at the Chalk
Point Steam Electric Station on the Patuxent River,
Maryland (Academy of Natural Sciences of Philadelphia

1983). The impingement of crabs and their subsequent
wash-off from the screens at the CCNPP had virtually no
affect on survival which exceeded 99% (Burton 1976).

Differences among years were detected for all population
variables examined and variation among stations over time
was moderate, but other than slightly larger males at
Kenwood Beach than at the other stations and a higher
percentage of males at Kenwood Beach than at Rocky
Point, no statistically significant station differences were
detected during pre-operational or operational periods.
Perhaps one of the most significant findings of this study,
however, was the long-term decrease in the percent of males
that occurred equally among stations. All year-to-year
changes in population structure, whether significant or not,
appeared to be natural fluctuations and unrelated to
operation of the CCNPP.

AC KNOWLEDGMENTS

1 thank all the individuals who assisted in the collection
of data during the 14 years of this project, but especially
Robert Cantin, Matt Newman, and William Yates, Jr. 1 am
also indebted to Elgin Perry for his computer analysis of
the data. This study was supported by the Baltimore Gas
and Electric Company.

REFERENCES CITED

Academy of Natural Sciences of Philadelphia , Marine Sciences
Research Center & J. E. Edinger Associates, Inc. 1980. Calvert
Cliffs Nuclear Power Plant thermal plume dye studies, April
and August 1979, and analysis of plume sites. Report No. 80-
10: 122 p. Available from: Academy of Natural Sciences,
Philadelphia, PA.

Academy of Natural Sciences of Philadelphia. 1983. Impingement.
Chalk Point 316 Demonstration of Thermal, Entrainment, and
Impingement Impacts on the Patuxent River in Accordance with
the Code of Maryland Regulation 08.05.04.13; 4:123-196.
Available from: Academy of Natural Sciences, Philadelphia, PA.

Baltimore Gas and Electric Company. 1970. Description of plant
effluent and waste systems. Environmental Report, Calvert Cliffs
Nuclear Power Plant. Pp. B1-B14. Available from: Baltimore
Gas and Electric Co., Baltimore, MD.

Burton, D. T. 1976. Impingement studies. II. Qualitative and quanti-
tative survival estimates of impinged fish and crabs. Semi-annual
Environmental Monitoring Report for the Calvert Cliffs Nuclear
Power Plant, March 1976. Pp 11.2-1-11.2-49. Available from:
Baltimore Gas and Electric Co., Baltimore, MD.

. 1978. The response of two estuarine Crustacea exposed to
time- temperature changes simulating once- through, 10 C AT,
power plant condenser entrainment. Rep. No. 78-30: 22 p.
Available from: Academy of Natural Sciences, Philadelphia, PA.

Cargo, D. G. 1960. A megalops of the blue crab, Callinectes sapidus,
in the Patuxent River, Maryland. Oiesapeake Sci. 1:110.

Costlow, J. D., Jr. & C. G. Bookhout. 1959. The larval development
of Callinectes sapidus Rathbun reared in the laboratory. Biol.
Bull. (Woods Hole) 116:373-396.

Cox, D. R. 1970. Analysis of Binary Data. London, U.K.: Chapman
and Hall. 142 p.

Gallaway, B. J. & K. Strawn. 1975. Seasonal abundance and distri-

bution of the blue crab, Callinectes sapidus Rathbun, in the dis-
charge area of the P. H. Robinson Generating Station, Galveston
Bay, Texas. Tex. J. Sci. 26:185-201.

Hicks, C. R. 1973. Fundamental Concepts in the Design of Experi-
ments. New York, NY: Holt, Rinehart and Winston. 349 p.

Hirshfield, M. F., J. H. Hixson, III & J. D. White. 1980. Impingement
studies. 1. Impingement counts. Nonradiological Environmental
Monitoring Report, Calvert Cliffs Nuclear Power Plant, January-
December 1979. Pp. 9.1-1-9.1-15. Available from: Baltimore
Gas and Electric Co., Baltimore, MD.

. 1981. Impingement studies. 1. Impingement counts. Non-
radiological Environmental Monitoring Report, Calvert Cliffs
Nuclear Power Plant, January-December 1980. Pp. 9.1-1-9.1-14.
Available from: Baltimore Gas and Electric Co., Baltimore, MD.

Hirshfield, M. F. & J. H. Hixson, III. 1982. Impingement studies.
1. Impingement counts. Nonradiological Environmental
Monitoring Report, Calvert Cliffs Nuclear Power Plant, January-
December 1981. Pp. 8.1-1-8.1-18. Available from: Baltimore
Gas and Electric Co., Baltimore, MD.

Lippson, A. J. 1973. 77?e Oiesapeake Bay in Maryland- an Atlas of
Natural Resources. Baltimore, MD: Johns Hopkins Univ. Press.
55 p.

May, E. B. 1973. Extensive oxygen depletion in Mobile Bay,
Alabama. Limnol. Oceanogr. 18:353-366.

Mihursky, J. A. & V. S. Kennedy. 1967. Water temperature criteria
to protect aquatic life. A Symposium on Water Quality Criteria
to Protect Aquatic Life. Amer. Fish. Soc. Spec. Publ. No. 4:
2-32.

National Marine Fisheries Service. 1972-1979a. Maryland landings,
1970-1978. Current Fisheries Statistics No. 5719, 5914, 6115,
6414, 6714, 6914, 7214, 7512 & 7717. U. S. Dept. of Commerce,
Washington, D.C.

Blue Crab Populations in Mid-Chesapeake Bay

193

. 1972-1979b. Virginia landings, 1970-1978. Current
Fisheries Statistics No. 5720, 5915, 6116, 6415, 6715. 6915.
7215, 7513 & 7718. U.S. Dept. of Commerce, Washington. D.C.
. 1980. Maryland landings, 1979. Current Fisheries Statistics

No. 8014. U.S. Dept. of Commerce, Washington, D.C.

_. 1981. Virginia landings, 1979. Current Fisheries Statistics
No. 8015. U.S. Dept. of Commerce, Washington, D.C.

. 1982. Preliminary commercial fishery landings, by state

(Maryland and Virginia). U.S. Dept. Comm., Natl. Mar. Fish.

Serv., Resour. Stat. Div., Washington, D.C.
Newcombe, C. L. 1945. The biology and conservation of the blue

crab, Callinectes sapidus Rathbun. Va. Fish. Lab. Educ. Ser.

No. 4: 39 p.
, F. Campbell & A. M. Eckstine. 1949. A study of the form

and growth of the blue crab Callinectes sapidus Rathbun. Growth

13:71-96.
Pearson. J. C. 1948. Fluctuations in the abundance of the blue crab

in Chesapeake Bay. U.S. Fish Wildl. Serv. Res. Rep. 14: 26 p.
Pullen, E. J. & W. L. Trent. 1970. Carapace width-total weight

relation of blue crabs from Galveston Bay, Texas. Trans. Am.

Fish. Soc. 99:795-798.
Sandoz, M. & R. Rogers. 1944. The effect of environmental factors

on hatching, moulting, and survival of zoea larvae of the blue

crab Callinectes sapidus Rathbun. Ecology 25:216-228.
Tagatz, M. E. 1965. The fishery for blue crabs in the St. Johns

River, Florida, with special reference to fluctuation in yield

between 1961 and 1962. U.S. Fish Wildl. Serv. Spec. Sci. Rep.,

Fish. 501: 11 p.

. 1969. Some relations of temperature acclimation and salinity

to thermal tolerance of the blue crab, Callinectes sapidus. Trans.

Am. Fish. Soc. 98:713-716.
Thoni, H. 1967. Transformations of variables used in the analysis

of experimental and observational data. A review. Ames, IA:

Iowa State Univ. Statistical Lab. Tech. Rep. No. 7 : 6 1 p.
Truitt, R. V. 1939. Our water resources and their conservation.

Solomons, MD: Chesapeake Biol. Lab., Contrib. No. 27: 103 p.
U.S. Fish and Wildlife Service. 1970a. Maryland landings, 1969.

U.S. Nat. Mar. Fish. Serv. Curr. Fish. Stat. No. 5307.
. 1970b. Virginia landings. 1969. U.S. Nat. Mar. Fish. Serv.

Curr. Fish. Stat. No. 5326.
Van Engel, W. A. 1958. The blue crab and its fishery in Chesapeake

Bay. I. Reproduction, early development, growth, and migration.

Commer. Fish. Rev. 20(6) :6- 17.

, 1962. The blue crab and its fishery in Chesapeake Bay.

II. Types of gear for hard crab fishing. Commer. Fish. Rev.

24(9):1-10.

Journal of Shellfish Research. Vol. 3, No. 2, 195-201, 1983.

MOVEMENTS OF TAGGED MALES OF TANNER CRAB

CHIONOECETES BAIRDI RATHBUN OFF

KODIAK ISLAND, ALASKA

WILLIAM E. DONALDSON

Alaska Department of Fish and Game
333 Raspberry Road
Anchorage, Alaska 99502

ABSTRACT From 1973 through 1978, 11,196 males of the Tanner crab Chionoecetes bairdi Rathbun were tagged and
released off of Kodiak Island. Alaska. A total of 1,961 tags was returned, 1,404 with accurate recovery data. Males which
were tagged in bays tended to move into offshore areas while those tagged offshore remained in that general area. Crab
movements were not extensive; mean net movement for all recoveries was 24 km (15 miles). The generalized movement
models indicate the presence of stocks of large male Tanner crabs in the Shelikof, Marmot-Chiniak, Eastside, and South-
west areas of Kodiak Island.

KEY WORDS Tanner crabs. Chionoecetes bairdi. migration, tagging, movement

INTRODUCTION

The Tanner crab Chionoecetes bairdi Rathbun occurs
from shallow nearshore areas to depths of 473 m (259 fm)
and ranges from Puget Sound, Washington (Slipp 1952) and
the Oregon coast (Hosie 1974) to the Aleutian Islands and
southeastern Bering Sea (Garth 1958) where male Tanners
are the basis for a major fishery (Otto 1981).

Many fishermen hold traditional beliefs concerning
Tanner crab migrations and cite time-related changes in
catch with depth as evidence of inshore-offshore movement.
Prior to this study, movement patterns of C. bairdi were
unknown; however, some information on migrations of
congeneric species does exist. Migration of the snow crab
C. opilio (O. fabricius) was studied in the Atlantic around
the Gaspe region of the Gulf of Saint Lawrence by Watson
(1970) and Watson and Wells (1972). Their results indicated
that tagged males traveled relatively little, with 85% of the
returns recaptured within 20.3 km ( 1 1 mi) of the release
points. Katoh et al. (1956) and Yoshida (1941) observed
bathymetric separation of the sexes of C. opilio in the Sea
of Japan indicating at least a seasonal migration for mating.
Pereyra (1967) concluded that males of C. tanneri off the
coast of Oregon showed seasonal variations in relative
abundance with depth, whereas the female population was
fairly stationary during all seasons, thus suggesting move-
ment of males for reproductive purposes.

In recent years, the fishery for C. bairdi has developed
exponentially, but data on the life history of this species
have not been accumulated in like manner. While C. bairdi
has accounted for about one fourth of the recent domestic
harvest of crabs by U.S. fishermen (Donaldson 1980),
resource data are insufficient to define discrete stocks in
most areas. The purpose of this study was to determine
whether migrations or displacement of aggregations of
males of C. bairdi occur over the shelf region surrounding
Kodiak Island.

MATERIALS AND METHODS

Migration was studied by the release and recapture of
tagged male crabs during a 6-year period (1973-1979).
Males of > 110 mm carapace width (CW) were tagged and
released between July 1973 and August 1975 off Kodiak
Island, AK. In 1976, minimum tagging size was raised to
135 mm CW because of the establishment of a commercial,
minimum size limit.

Crabs were tagged with a combination of Floy disc,
FD 67 "T" bar, and a modified FD 67 "T" bar (also known
as the McBride tag). The Floy disc is a temporary tag which
is lost during ecdysis. The FD 67 "T" bar and modified
FD 67 "T" bar are prototype permanent tags. Floy disc
tags were used in all years except 1977; both FD 67 "T"
bars and Floy discs were used in 1975; and only the modi-
fied 67 "T" bar was used in 1 977.

Crabs were captured with 2.1- X 2.1-m (7- X 7-ft) crab
pots which were covered with 89-mm (3.5-in.) mesh. Tag
number, date, location, and depth of capture were recorded
for crabs tagged from 1973 through 1975. Exoskeletal age
(intermolt period) and carapace width were also recorded
beginning in 1976. Females, generally, are too small to
be captured in pots and none were tagged during this study.

Crabs were captured, tagged, and released at various
inshore (bay) and offshore locations (Figure 1). Tagged
crabs were recovered from fishermen and at processing
plants. Recovery data included tag number, date, location
and depth of capture, carapace width, and exoskeletal age.
Only recaptures with complete recovery data were used in
this study.

Distance and direction of net migration and the absolute
depth change were recorded for the year of release and
for inshore (bay) and offshore areas. All recovery data
were collated by specific geographical area for the duration
of the study and migration by specific areas was analyzed.
The data were insufficient to assess migration by cohorts;

195

196

Donaldson

— 59°N

such group was determined. Changes in depth were deter-
mined and represent the percentage of crabs that were
recovered deeper, shallower, or remained at the release
depth. Directions of movement and recovery locations
were analyzed using computer-calculated ellipses that
represented a 95% confidence interval of direction and
recovery region data. Information on the distribution of
fishing effort was obtained from a fish-ticket reporting
system.

— 59°N

156°

155

Figure 1. Release sites of males of the Tanner crab Chionoecetes
bairdi Rathbun at Kodiak, AK. Top: bay sites (7) 1973-1978.
Middle: offshore sites (X) 1973-1975. Bottom: offshore sites (X)
1976-1978. Alitak Bay (A), Chiniak Bay (C), Eastside area (E),
Kiliuda Bay (K), Kupreanof-Viekoda Bay area (K-V), Marmot-
Chiniak Bay area (M-C), Marmot-Kizhuyak Bay area (M-K),
Sitkalidak Bay (S), Shelikof area (SH), Southwest area (SW).

therefore, the crabs were grouped by 30-day periods
between release and recovery. The mean movement by each

RESULTS

1973

A total of 2,285 male crabs (> 1 10 mm CW) were tagged
and released between 15 July and 5 August (Table 1 ). The
majority (2,024) was released in offshore areas, while
261 tagged crabs were released in inshore bays. A total of
486 recoveries were made, 415 from offshore and 71 from
inshore (bays). Catch data were available from 361 (15.8%)
of the tagged crabs. Time of freedom ranged from 26 to
1,376 days (Table 2). Crabs recovered within one, two, and
three years of release represented 72.5%, 19.1%, and 7.4%
of total recoveries, respectively. Three crabs (0.8%) were
captured in their fourth year after tagging. Mean migration
distance (based on two or more recovered crabs) ranged
from 49.6 km (30.8 mi) for nine crabs that were free
between 961 and 990 days, to 11.5 km (7.1 mi) for two
crabs that were free 481 to 510 days (Table 2). The mean
absolute distance traveled was 27.9 km (17.3 mi) for all
recovered crabs. Crabs that were tagged and released in
bays tended to move offshore while those tagged and
released offshore remained offshore and within the
geographic area of release. Depth change was variable, with
175 crabs (48.4%) recaptured at points shallower than
their release depth, 179 (49.6%) recaptured at deeper
points, and 7 ( 1 .9%) recaptured at their release depth.

1974

During 1974, 1,846 male crabs ( > 1 10 mm CW) were
tagged with Floy disc tags and released (Table 1). The
majority (1,472) was released in offshore areas, while 374
were released in bay areas. A total of 397 tags were
recovered (340 offshore, 57 inshore). Catch data were
available from 310 recoveries or 16.8% of all crabs tagged.
Time of freedom ranged from 30 to 1.080 days (Table 2).
Crabs recaptured within one, two, and three years of release
represented 57.7%. 36.9% and 5.4% of all recoveries,
respectively. The longest mean (absolute) movement was
83.5 km (51.9 mi) for four crabs that were free between
991 and 1,020 days; the shortest mean movement was
12.0 km (7.5 mi) for two crabs that were free from 1,021
to 1,050 days (Table 2). The overall mean distance of net
migration was 26.8 km (16.7 mi). Movement in all offshore
areas was localized within the area of release. Crabs tagged
and released in bays tended to move offshore as did the

Movements of Tagged Tanner Crabs

197

I
s

c
H

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z£^

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198

Donaldson

TABLE 2.

Distance moved from release site for males of the Tanner crab Chionoecetes bairdi Rathbun off Kodiak Island, AK, 1973-
Mean movement from release site in km and number of crabs indicated within parentheses.

1978.

Days of
Freedom

1975*

1973

1974

ADFG

NMFS

1976

1977

1978

All Years

1- 30

31- 60

61- 90

91- 120

121- 150

151- 180

181- 210

211- 240

241- 270

271- 300

301- 330

331- 360

One Year

361- 390
391- 420
421- 450
451- 480
481- 510
511- 540
541- 570
571- 600
601- 630
631- 660
661- 690
691- 720

Two

721-
751-
781-
811-
841-
871-
901-
931-
961-
991-
1021-
1051-

Years

- 750

- 780
810
840

- 870
900
930

- 960
990

1020
1050
1080

Three Years
1343-1376

23.7
24.0
11.0
31.6
35.3
17.9
26.9
30.8
27.9
22.3

24.7
12.8

11.5
56.0

31.7
23.2
30.8
13.0

31.7
49.0
29.0

17.0
23.5
49.6
43.1
38.5

34.3

(3)

(1)

(23)

(12)

(21)

(39)

(106)

(33)

(0)

(0)
(8)
(4)
(0)
(2)
(1)
(0)
(0)
(3)
(29)
(21)
(1)

(0)
(0)
(3)
(1)
(2)
(0)

(1)

(2)
(9)
(7)
(2)
(0)

(3)

17.1
24.8
22.7
25.0
60.0

43.3
28.6
19.1
14.3

25.2
19.6
24.0
38.5
20.3
24.8
24.0
24.3

19.0

41.3
29.5
83.5
12.0
30.0

(1)

(17)

(6)

(1)

(0)

(24)

(78)

(48)

(4)

(0)

(4)

(5)

(2)

(14)

(19)

(12)

(25)

(34)

(0)

(0)
(0)
(0)
(1)
(0)
(0)
(0)
(3)
(4)
(4)
(2)
(1)

(0)

7.3
8.0

15.4
19.4
17.4
16.0
12.3
20.6

4.0
32.0

20.5
25.4
20.5
41.0

138.0
55.0

(0)

(3)

(1)

(0)

(22)

(9)

(20)

(26)

(19)

(5)

(0)

(0)
(1)
(1)
(0)
(0)
(0)
(4)
(8)
(2)
(1)
(0)
(0)

(0)
(0)
(0)
(0)
(0)
(0)
(1)
(1)
(0)
(0)
(0)
(0)

(0)

30.0
38.1
25.5
14.9
15.8

(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)
(0)

(0)
(1)
(2)
(30)
(36)
(5)
(0)
(0)
(0)
(0)
(0)
(0)

(0)
(0)
(4)
(0)
(2)
(0)
(0)
(0)
(0)
(0)
(0)
(0)

(0)

21.0

15.0

17.5
18.4
18.5
23.0
25.8
46.0

10.0

14.0
39.5
9.3
8.5
26.0
29.0
23.0

31.0

41.0
28.5
34.0

(0)

(9)

(46)

(57)

(59)

(38)

(1)

(0)

(1)

(0)

(2)

(3)

(1)

(11)

(60)

(21)

(0)

(0)
(0)
(0)
(0)
(0)
(0)
(1)
(3)
(2)
(1)
(0)
(0)

11.4
9.8
9.7
32.3
15.3
17.7
15.9
24.0

47.0

40.5
20.0
25.5
20.7

(0)

(2)

(5)

(7)

(4)

(11)

(38)

(34)

(47)

(0)

(0)
(0)
(1)
(0)
(0)
(2)
(2)
(2)
(3)
(0)
(0)
(0)

(0)

6.0
87.0

19.0
19.5
17.3
18.4

(0)
(1)
(1)
(0)
(0)
(1)
(4)
(27)
(19)
(0)
(0)
(0)

(0)

23.5
20.9
20.8
26.0
24.4
17.2
19.5
20.1
25.8
26.3
19.6
14.3

10.0
23.2
28.8
24.1
17.7
25.7
19.9
25.4
26.8
23.8
30.8
13.0

25.6
34.0
19.5

62.0
38.8

41.4
55.8
25.3
30.0

(4)

(26)

(14)

(31)

(39)

(51)

(129)

(183)

(274)

(154)

(49)

(4)

(1)
(10)
(12)
(37)
(42)
(25)
(26)
(33)
(93)
(85)
(21)

(1)

(0)
(0)
(7)
(2)
(4)
(0)
(3)
(9)
(15)
(12)
(4)
(1)

361

310

124

318

158

1404

2km/N

= mean (km)

27.9

26.8

18.2

21.1

23.1

19.4

19.0

24.0

*ADFG. Alaska Department of Fish and Game; NMFS. National Marine Fisheries Service.

1973 releases. Of the 310 returns, 168 (54.2%) were
recovered at points deeper than their release depth, 108
(34.8%) were recovered at shallower points, and 34 (10.9%)
were recaptured at their release depth.

1975

Between 15 July and 12 November 1975, 2,106 crabs

1,174 were tagged with FD 67 "T" bar tags and 932 crabs
bore Floy discs. A total of 1,268 (60.2%) crabs were
released in offshore areas while 838 (39.8%) were released
in bays (inshore). A total of 325 were recovered, 226 from
offshore and 99 from inshore areas. Catch data are available
from 204 of the recoveries or 9.7% of all crabs tagged.
Time of freedom ranged from 40 to 935 days (Table 2).

(3 s 1 10 mm CW) were tagged and released (Table l);of these. Crabs recovered within one, two and three years of release

Movements of Tagged Tanner Crabs

199

accounted for 48.8%, 47.4%, and 3.8% of all recoveries,
respectively. The longest mean net movement was 38.1 km
(23.6 mi) for two crabs that were free from 421 to 450 days
(Table 2). The shortest mean movement was 7.3 km (4.5 mi)
for three crabs that were free from 31 to 60 days. The
overall mean distance of migration was 19.3 km (12.0 mi).
Little difference existed in the mean (absolute) distances of
migration for crabs tagged with the FD 67 "T" bar (21.1 km
[13.1 mi]) versus those bearing Floy discs (18.2 km [ 1 1 .3
mi] ). Recoveries were restricted to the northeastern and
eastern sides of the island. Movement of offshore crabs was
localized around the area of release while crabs tagged
inshore moved offshore. Forty-one (20.1%) crabs were
recaptured at points shallower than their release, 153
(75.0%) were recaptured at deeper points, and 10 (4.9%)
were recaptured at their release depth.

1976

Between 24 June and 8 August 1976, 2.324 crabs
(> 135 mm CW) were released bearing Floy disc tags. As
in previous years, the majority of tagged crabs (2,023
[87%]) were released in offshore areas; 301 (13%) were
tagged and released inshore (bays) (Table 1). A total of
499 recoveries were made of which 434 were crabs tagged
and released offshore and 65 were crabs that had been tagged
and released inshore. Catch data are available from 318
recoveries or 13.7% of the total releases. Time of freedom
ranged from 166 to 994 days (Table 2). Crabs recovered
within one, two, and three years of release represented
66.3%. 31.5% and 2.2%, respectively, of total recoveries.
Mean movement by 30-day periods ranged from 9.3 km
(5.8 mi) for three crabs that were free from 511 to 540 days
to 41.0 km (25.5 mi) for three crabs that were free from
931 to 960 days. The mean (absolute) distance traveled by
all 318 crabs was 23.1 km (14.4 mi) (Table 2). Two-
hundred nine crabs (65.7%) were recovered from points
shallower than release depth and 109(34.3%) were recovered
from water deeper than release depth.

1977

Between 27 June and 18 August 1977, 1,672 crabs
(> 135 mm CW) were tagged with the modified FD 67 "T"
bar tag; 1,351 (80.8%) were released in offshore areas,
while 321 (19.2%) were released in bays. A total of 181 tags
were recovered (167 offshore, 14 inshore). Catch data are
available from 158 or 9.4% of the total releases. Time of
freedom ranged from 44 to 61 7 days (Table 2). The majority
of the recoveries (148 [93.7%] ) occurred within one year
of release, the remainder ( 10 [6.3%] ) were recovered during
the second year. Mean movement ranged from 9.7 km
(6.0 mi) for seven crabs that were free from 91 to 120 days
to 40.5 km (25.2 mi) for two crabs that were free 511 to
540 days. The mean (absolute) distance migrated for all
158 crabs was 19.4 km (12.1 mi) (Table 2). Movement
patterns were consistent with those from previous years

and regions where tagging took place. The majority of crabs
(105 [66.5%]) were recovered from points shallower than
their release sites, 43 (27.2%) had moved deeper, while
10 (6.3%) were recaptured at their release depth.

1978

A total of 963 crabs (> 135 mm CW) were tagged with
Floy disc tags and released in the southern portion of the
Kodiak Island area. No tagging was done in bays. Seventy-
three recoveries were made; reliable catch data are available
for 53 or 5.5% of the total releases. Time of freedom ranged
from 55 to 263 days (Table 2). The longest mean (absolute)
movement was 19.5 km (1 1 .8 mi) (Table 2). No crabs tagged
offshore were recovered inshore. Thirty-seven crabs (71 .2%)
were recovered in depths shallower than release depths,
15 (28.8%) were recovered at deeper depths, and 1 crab was
recaptured at the same depth as released.

Inshore Areas, 1973-1978

Tagging and recovery trends for inshore (bay) areas for
all years combined are depicted in Figure 2. A total of
212 tagged crabs with recovery data from all bays (Table 1 )
were obtained during the course of this study; they ranged
from 52 crabs from Sitkalidak Bay (S) to 17 from the
Marmot-Kizhuyak Bay area (M-K). Movement from the
midpoint of the release locations to the midpoint of the
recovery locations was greatest in Marmot-Kizhuyak Bay
(33.6 km [20.9 mi] ), while the least movement occurred
in theKupreanof-ViekodaBay(K-V) area (9.6 km [6.0 mi] ).
Movement of crabs tagged and released in Kiliuda (K) and
Alitak (A) bays averaged 14.4 km (8.9 mi);Chiniak Bay (C)
movement averaged 17.6 km (10.9 mi); and Sitkalidak Bay
on the southeastern side of Kodiak Island averaged 22.3 km
(13.9 mi). All crabs tagged and released in bays demonstrated
an offshore movement with the exception of Kupreanof-
Viekoda Bay area recoveries, which demonstrated both
offshore and onshore movement.

Offshore Areas, 1973-1978

Of the tagged crabs released in offshore areas from 1973
to 1978, 1,192 were recovered with complete catch data
(Table 1 ). Direction and magnitude of migration are depicted
in Figure 2. Four individual stocks are somewhat apparent
on that figure: (1) Marmot-Chiniak (M-C) area, 230 taggs
recovered; (2) Eastside (E), 494 tags recovered; (3) Southwest
(SW), 453 tags recovered; and (4) Shelikof (SH), 15 tags
recovered. Crabs appeared to move around in the area of
tagging and release with no immigration into bay areas or
adjacent stocks. There is an apparent westerly movement
of crabs released in the northern portion of the Eastside
and Shelikof areas; however, because of the small number
of tagged crabs recovered, the data do not permit a firm con-
clusion. The apparent northerly movement in the southern
portion of the Eastside area was probably caused by a lack
of commercial fishing to the south of the release points.

200

Donaldson

The Marmot-Chiniak and Eastside stocks are separated
by a deep gully of 144 to 215 m (80 to 120 fm) depth;
that gully may be a physical barrier that separates postlarval
crabs into independent stocks. Likewise, the Eastside and
Southwest stocks are separated by a large shallow area of
18 to 36 m (10 to 20 fm) running northeast-southwest;
that ridge may also limit or cross channel movement.

59° N

— 58°

-57°

— 56°

156° 155° 154'

53° 152° W

59° N

156°

155°

154°

152° W

Figure 2. Movements of tagged males of the Tanner crab Chionoecetes
bairdi Rathbun released from 1973 through 1978 at Kodiak, AK.
Top: inshore (bay) recoveries (212). Bottom: offshore recoveries
(1,192). (Ellipse represents 95% confidence region.) Alitak Bay (A),
Chiniak Bay (C), Eastside area (E), Kiliuda Bay (K), Kupreanof-
Viekoda Bay area (K-V), Marmot-Chiniak Bay area (M-C), Marmot-
Kizhuyak Bay area (M-K), Sitkalidak Bay (S), Shelikof area (SH),
Southwest area (SW).

DISCUSSION

Tag recovery is dependent on when and where fishermen
place their crab pots. From 1973 through 1978, 57,334.7 mt
(124,809,323 lb) of Tanner crabs were harvested off
Kodiak Island. Those landings represented approximately
49,934,730 crabs at 1.13 kg (2.5 lb) per crab. After release,
tagged crags were first subjected to recapture in the fall
fishery (August- December) for the king crab Paralithodes
camtschatica (Tilesuis). (Tanner crabs are captured
incidental to king crabs because the two species tend to
share the same habitat.) Tagged crabs were then subjected
to recapture during the Tanner crab fishery that opens
between November and January and closes m April or
May.

From 1973 to 1978 fishing effort expanded to cover all
major habitats of king and Tanner crabs. Fishermen with
smaller vessels tended to fish the nearshore areas while
fishermen with larger vessels primarily fished the deeper,
offshore areas. The tag-recovery data were influenced by
the peculiarities in fishing patterns; however, recovery of
tagged crabs appeared to be reasonably well distributed
over the study area and should provide a reasonable picture
of migration.

Tagged males did not move extensively from their release
sites. The results of this study demonstrated that although
there were examples of extensive movement for small
numbers of crabs, the mean (absolute) movement was only
24.0 km (15.0 mi). Although periods of freedom for tagged
individuals varied from less than one month to 3.8 years, no
correlation between time and absolute distance migrated
was evident. Watson (1970) and Watson and Wells (1972)
demonstrated a mean movement of 20.3 km (11 mi) for
adult males of Chionoecetes opilio. Male Tanner crabs that
were captured and tagged in bay areas tended to move to
deeper, offshore waters while those captured and tagged in
offshore waters remained offshore and migrated randomly
within a geographic area. These findings have implications
for management of the resource. High exploitation rates
in offshore areas may be partially compensated for by
immigration of mature crabs from bays. High exploitation
rates in bays may present a more difficult management
situation because recruitment into the fishable size range
is dependent on annual recruits to legal size with no
apparent immigration of offshore crabs.

An additional result of this study is that postlarval crabs
may be separated into manageable stocks because there is
little or no apparent movement between designated
geographic regions. Additional tag-and-recapture studies in
the vicinity of apparent geographic stock boundaries and
bathymetric features should help demonstrate whether or
not those apparent stocks are distinct or an artifact of
aggregated release locations.

Movements of Tagged Tanner Crabs

201

ACKNOWLEDGMENTS I thank Dr. Jerry Reeves of the Montlake Laboratory of

I acknowledge Matthew Dick, David Hicks, Rich Peterson, the National Marine Fisheries Service, Seattle, WA, for the
Mary Clemens, and Marilyn Kemerer for their contributions, use of his crab tagging data.

references cited

Donaldson, W. E. 1980. Alaska Tanner crab investigations. Alaska
Dep. Fish GameComp. Rep. Prof. No. 5-41-R: 122 p.

Garth, J. S. 1958. Brachyura of the Pacific Coast of North America.
Oxyrhyncha. Allen Hancock Pacific Exped. Los Angeles, CA:
University Southern California Press. 21: 854 p.

Hosie. M. J. 1974. Southern range extension of the Baird crab.
Chionoecetes bairdi Rathbun. Calif. Dep. Fish Game Fish Bull.
60:44-47.

Katoh, G., 1. Yamanaka, A. Ochi & T. Ogata. 1956. General aspects
on trawl fisheries in the Japan Sea. Bull. Jpn. Sea Reg. Fish.
Res. Lab. 4:1-331. (In Japanese with English summary; transla-
tion of pp. 293-305 available from U.S. Natl. Mar. Fish. Serv.
Trans. Prog., Seattle, WA.)

Otto, R. S. 1981. Eastern Bering Sea crab fisheries. Wood, D. W. and
J. A. Calder, eds. Tlie Eastern Bering Sea Shelf: Oceanography
and Resources. Seattle, WA: Univ. Washington Press. Vol. II:
1037^1066.

Pereyra, W. 1967. The bathymetric and seasonal abundance and

general ecology of the Tanner crab, Chionoecetes tanneri Rathbun
(Brachyura: Majidae), off the northern Oregon coast. Seattle,
WA: Univ. Washington. Thesis. 415 p.

Rathbun. M. J. 1924. The spider crabs of America. U.S. Natl. Mus.
Bull. 129:613 p.

Slipp, J. W. 1952. Status of crab, Oiionoecetes bairdi, in the inshore
waters of Washington and British Columbia. Wasmann J. Biol.
10:235-239.

Watson, J. 1970. Tag recaptures and movements of adult male snow-
crabs. Oiionoecetes opilio (O. fabricius) in the Gaspe region of the
Gulf of S t. Lawrence. Fish. Res. Board Can. Tech. Rep. No. 204:1 6p.

& P. G. Wells. 1972. Recaptures and movements of tagged

crabs (Oiionoecetes opilio) in 1970from theGulfof St. Lawrence.
Fish. Res. Board Can. Tech. Rep. No. 349:12 p.

Yoshida, H. 1941. On the reproduction of useful crabs in North
Korea (II). Suisan Kenkyushi 36:116-123. (In Japanese; transla-
tion of pp. 116-121 available from U.S. Natl. Mar. Fish. Serv.
Trans. Prog., Seattle, WA.)

Journal of Shellfish Research. Vol. 3, No. 2, 203-205, 1983.

RESEARCH NOTE

CHEMICAL INDUCTION OF SPAWNING BY SEROTONIN IN THE
OCEAN QUAHOG ARCTICA ISLANDICA (LINNE)

M. C. GIBBONS; J. G. GOODSELLt M. CASTAGNA 1
AND R. A. LUTZ 2

1 Virginia Institute of Marine Science and
School of Marine Science

College of William and Mary
Wachapreague, Virginia 23480

2 Department of Oyster Culture

New Jersey Agricultural Experiment Station
Cook College, Rutgers University
New Brunswick, New Jersey 08903

ABSTRACT Serotonin injected into the anterior adductor muscle induced spawning in the ocean quahog Arctica
islandica (Linne) when using either individual or mass spawning techniques. This represents the first successful attempt to
induce the release of gametes in this species which historically has been unresponsive to conventional spawning stimuli. The
gametes released were competent and fertilization occurred without treating the encapsulated eggs with ammonium
hydroxide or other chemicals. Larvae were reared through metamorphosis to early juvenile stage.

KEY WORDS: Ocean quahog, Arctica islandica, spawning, serotonin

INTRODUCTION

The ocean quahog .4 re /7a7 islandica (Linne) spawns from
August through November on the southern New England
shelf and off New Jersey (Jones 1981, Mann 1982). Attempts
to spawn the ocean quahog in the laboratory have been
unsuccessful. Various combinations of stimuli such as
thermal shock, addition of gonadal products, salinity and
pH changes, and exposure to hydrogen peroxide, which
are effective with many other bivalve species, have not
induced spawning (Loosanoff 1953, Landers 1976, Lutz
et al. 1 982. Mann 1 982). All larvae of ocean quahogs cultured
to date under laboratory conditions have been reared from
stripped gametes that had been fertilized after pretreatment
of eggs with ammonium hydroxide (Landers 1976, Lutz
et al. 1981).

Serotonin (5-hydroxytryptamine, creatinine sulfate
complex) has proven to be an effective chemical inducer of
spawning for many bivalve species (Matsutani and Nomura
1982, Gibbons and Castagna [in press]). The injection of
serotonin into the anterior adductor muscle or gonad of
certain bivalve species when ripe will induce spawning
using individual spawning techniques without any additional
stimuli. The present study describes the successful spawning
of ocean quahogs in the laboratory using serotonin.

MATERIALS AND METHODS

Sexually mature ocean quahogs, ranging in shell length
from 8 to 13 cm, were obtained in October 1983 using a

Contribution No. 1220 from Virginia Institute of Marine Science.
Publication No. D-32401-2-85, supported by state and various
National Oceanic and Atmospheric Administration Sea Grant funds
to Rutgers University.

commercial hydraulic dredge in 50 to 80 m of water off
Cape May, NJ. The specimens were kept on ice for approxi-
mately 1 2 hours during transport from the sampling site.
Upon arrival at the Wachapreague Laboratory of the Virginia
Institute of Marine Science, half of the ocean quahogs
were immediately placed in individual dishes of seawater
for spawning while the other half were held in a recirculating
seawater table at 15— 16°C.

A 2-mM solution of serotonin (Sigma Chemical Company,
St. Louis, MO) was prepared by dissolving crystalline
serotonin in l-/im-filtered seawater. Each ocean quahog was
washed and a small notch filed into the valve margin
adjacent to the anterior adductor muscle. To induce
spawning, 0.4 mC of the 2-mM serotonin solution was
hypodermically injected into the anterior adductor muscle.

Both individual and mass spawning techniques as
described by Castagna and Kraeuter (1981) were utilized
without any thermal shock or other stimulation to spawn
ocean quahogs. All spawning experiments were conducted
at a salinity of 32 ppt and at a controlled temperature of
15-16°C. Ocean quahogs were spawned by placing single
specimens in glass dishes containing IS of l-/jm-filtered
seawater. Mass spawning was achieved by placing the
quahogs in troughs containing 140 2 of static, l-/jm-filtered
seawater. Equal numbers of quahogs in the control groups
were treated in the same manner as the test groups except
they were injected with 0.4 m? of l-/am-filtered seawater
instead of the serotonin solution. The control animals from
trial 1 of the mass spawning were the test group for trial 2.
The G-test of independence and Williams' correction for a
2X2 contingency table were used to statistically determine

203

204

Gibbons, et al.

whether spawning was independent of injection with the
serotonin solution (Sokal and Rohlf 1981).

Eggs obtained from the serotonin-induced spawnings were
fertilized using standard techniques developed for other
bivalves (Loosanoff and Davis 1963, Castagna and Kraeuter
1981). Eggs were not pretreated with ammonium hydroxide
or other chemicals prior to fertilization. The larvae were
reared through settlement and metamorphosis to early
post-set at 13.5°C.

RESULTS AND DISCUSSION

Injection of the serotonin solution induced gamete
release in both the individual and mass spawning trials,
although greater percentages (35.5% and 37.1%) of ocean
quahogs spawned using the mass spawning technique than
for the individual method (17.1% and 22.5%) (Table 1 ). In
each case larger numbers of quahog males spawned than
females. This, however, may be a dose response. Ocean
quahogs injected with serotonin extended their siphons,
probed with their feet, and began spawning within
15 minutes. The control groups injected with filtered sea-
water did not exhibit any of these behavioral patterns and
did not spawn.

The egg capsules of the ocean quahog are unlike any
structures described for bivalves (Castagna et al. 1982). The
encapsulated eggs were slightly ovoid and ranged from
75.0 to 85.0 jum in diameter (X = 79.9 jim;S.D. = 1.3 Mm).
Fertilization occurred in mass spawnings and similarly upon
addition of sperm in individual spawnings without chemical
pretreatment of the freshly spawned eggs. The egg capsules
have been suggested as being responsible for the difficulty
in spawning ripe ocean quahogs or in fertilizing stripped
eggs (Lutz et al. 1982). but no difficulty was observed with
this technique. Exposure of stripped eggs to ammonium

hydroxide may result in a lower percentage of normally
developing larvae compared to naturally spawned eggs
(Loosanoff and Davis 1963). Serotonin-induced spawning
appears to be a more effective means of obtaining gametes
from ripe ocean quahogs than stripping gametes from
mature individuals.

The development of larvae from the trochophore stage
through metamorphosis was similar to that described for
larvae of this species obtained from fertilization of stripped
eggs (Landers 1976; Lutz et al. 1981, 1982). Developing
eggs were encapsulated up to the gastrula stage, at which
time the egg capsules were lost. Metamorphosis occurred
at shell lengths of 170.6 to 266.7 |um (X = 220.5 /im;
S.D. = 19.8 /im) between 37 and 62 days after natural
fertilization, which was similar to results obtained by others
for fertilized strippedeggs (Landers 1976, Lutz et al. 1982).

To date, serotonin has been effectively utilized to induce
spawning in several species of bivalves (Matsutani and
Nomura 1982. Gibbons and Castagna [in press]). It is a
neurotransmitter that occurs naturally in the cerebropleural,
pedal, and visceral ganglia of Arctica islandica at concentra-
tions of 20 jug • g fresh tissue" 1 (Welsh and Moorhead 1960).
In laboratory studies, serotonin has been found to excite
excised hearts of ocean quahogs by stimulating the cardio-
regulatory nerves (Gaddum and Paasonen 1955, Leake and
Walker 1980). The physiological role of serotonin as an
inducer of spawning in bivalves is unknown.

The use of serotonin has induced spawning in the ocean
quahog, a bivalve that historically has been difficult to
spawn in the laboratory. Serotonin has potential value to
induce spawning in other bivalves which are resistant to
conventional spawning stimuli. The advantages of this
technique include ease of use and rapid and synchronous
spawning of ripe individuals.

TABLE 1.

Numbers of ocean quahogs induced to spawn by injection of serotonin.

Spawning Technique

Treatment

Number Tested

Number Spawned

Percentage Spawned

Number Males

Number Females

Individual - trial 1

Serotonin

17.1

Control

Individual - trial 2

Serotonin

22.5

Control

Mass — trial 1

Serotonin

13*

37.1

Control

Mass - trial 2

Serotonin

35.5

'significant at P < 0.005.

Chemical Induction of Spawning by Serotonin

205

REFERENCES cited

Castagna. M., J. Goodsell. R. Lutz & R. Mann. 1982. The egg capsule

of Arctica islandica. J. Shellfish Res. 2:91-92.
Castagna, M. & J. N. Kraeuter. 1981. Manual for growing the hard

clam Mercenaria. Va. Inst. Mar. Sci. Spec. Rep. Appl. Mar. Sci.

Ocean Eng. 249: 110 p.
Gaddum, I. H. & M. K. Paasonen. 1955. The use of some molluscan

hearts for the estimation of 5-hydro\ytryptamine. Br. J.

Pharmacol. 10:474-483.
Gibbons, M. C. & M. Castagna. (in press) Serotonin as an inducer

of spawning in six bivalve species. Aquaailture .
Jones, D. S. 1981. Reproductive cycles of the Atlantic surf clam,

Spisula solidissima. and the ocean quahog, Arctica islandica.

off New Jersey. J. Shellfish Res. 1:23-32.
Landers, W. S. 1976. Reproduction and early development of the ocean

quahog, Arctica islandica, in the laboratory. Nautilus 90:88-92.
Leake. L. D. & R. J. Walker. 1980. Invertebrate Neuropharmacology .

New York, NY: John Wiley and Sons. 102-143.
Loosanoff, V. L. 1953. Reproductive cycle in Cyprina islandica.

Biol. Bull. (Woods Hole) 104:146-155.

& H. C. Davis. 1963. Rearing of bivalve mollusks. Adv.

Mar. Biol. 1:1-136.
Lutz, R. A., J. G. Goodsell, R. Mann & M. Castagna. 1981. Experi-
mental culture of the ocean quahog, Arctica islandica. J. World

Maricul. Soc. 12:196-205.
Lutz. R. A., R. Mann. J. G. GoodseU & M. Castagna. 1982. Larval

and early development of Arctica islandica. J. Mar. Biol. Assoc.

U.K. 62:745-769.
Mann, R. 1982. The seasonal cycle of gonadal development in

Arctica islandica from the Southern New England Shelf. U.S.

Natl. Mar. Fish. Serv. Fish. Bull. 80:315-326.
Matsutani, T. & T. Nomura. 1982. Induction of spawning by

serotonin in the scallop. Patinopecten yessoensis (Jay). Mar.

Biol. Lett. 3:353-358.
Sokal, R. R. & F. J. Rohlf. 1981. Biometry. 2nd ed. San Francisco,

CA : W. H. Freeman & Co. 859 p.
Welsh, J. H. & M. Moorhead. 1960. The quantitative distribution of

5-hydroxytryptamine in the invertebrates, especially in their

nervous systems. J. Neurochem. 6:146-169.

Journal of Shell fish Research, Vol 3, No. 2, 207-221, 1983.

NATIONAL SHELLFISHERIES ASSOCIATION

ACTIVE MEMBERS

(As of 1 January 1984)

*Denotes Honorary Members

ABBOTT, Dr. R. Tucker, American Malacologists, Inc., P.O. Box

2255, Melbourne, FL 32901
ADAMKEWICZ. Dr. S. Laura, Dept. of Biology, George Mason Univ.,

4400 University Drive, Fairfax, VA 22030
AKASHIGE, Satoru, Hiroshima Fisheries Experiment Station,

5233-2 Ondo, Aki-gun, Hiroshima 737-12 Japan
ALLEN, Donald. National Marine Fisheries Service, Southeast

Fisheries Center. 75 Virginia Beach Dr., Miami, FL 33149
ALLEN, Standish, K., 313 Murray Hall, Univ. of Maine, Orono,

ME 04469
ALATALO, Philip, Marine Biological Laboratory, Woods Hole,

MA 02543
ALPER1N, Irwin M., Atlantic States Marine Fisheries Commission,

1717 Massachusetts Ave., NW, Washington, DC 20036
ANDERSON, Bruce A., 105 A. Kelly Rd., Clemson, SC 29631
ANDERSON, W. C, South Carolina Marine Research Inst., P.O.

Box 12559, Charleston, SC 29412
'ANDREWS, Jay D , Virginia Institute of Marine Science, Gloucester

Point, VA 23062
APLIN, J. A., RR 4, Box 268W, Newport, NC 28570
APPELDOORN, Dr. Richard, Dept. of Marine Sciences, Univ. of

Puerto Rico, Mayaguez, PR 00708
APTS, Charles W.. Battelle Marine Research Lab., 439 West Sequim

Bay Rd., Sequim, WA 98382
ARAKAWA, Dr. Kohman Y.. Fishery Section/Hiroshima Prefectural

Government, 10-52 Moto-machi, Hiroshima 730. Japan
ARMSTRONG, Dr. David, School of Fisheries WH-10, Univ. of

Washington, Seattle, WA 98195
ARNOLD, Bill, Harbor Branch Inst. Inc.. RR No. 1, Box 196A,

Fort Pierce, FL 33450
ARY, Roy D.. Dept. of Biological Science, Univ. of New Orleans,

Lakefront, New Orleans, LA 70148
AUSTER, Peter, National Undersea Research Program, Univ. of

Connecticut, Groton, CT 06340

BACON, Dr. G. B., Research & Productivity Council, Box 6000,

Fredericton, New Brunswick. Canada E3B 5H1
BAGLIN, Raymond E., P.O. Box 2969, Kodiak, AK 99615
BAQUEIRO, Erik, Apartado Postal 46B, La Paz, Baja California,

Mexico
BARBER, Bruce J., Dept. of Marine Science, Univ. of South Florida,

140 7th Ave., St. Petersburg, FL 33701
BARCELLOS, Lauro, Museu Oceanografico, P.O. Box 379, Rio

Grande 96200 R.S., Brasil
BARRY, Steven T., Washington Dept. of Fisheries, 331 State High-
way 12, Montesano, WA 98563
BASS, Ann E„ 94 Neal Street, Portland, ME 04102
BAYER, Dr. Robert, Dept. of Animal Vet. Science/Hitchner Hall,

Univ. of Maine, Orono, ME 04469
BEAL, Brian F., Univ. of Maine at Orono. Cooperative Extension

Service, 5 Cooper St., Machias, ME 04654
BEATTIE, J. Harold, National Marine Fisheries Service Aquaculture

Station, P.O. Box 38, Manchester, WA 98353
BENNETT, Dr. Joseph T., Dept. of Chemistry, Bowdoin College,

Brunswick, ME 04011
BENNETT, Leonard, R & B Oyster, Inc., Box 321. Bay Center,

WA 98527

BERR1GAN, Mark E., Dept. of Nat. Resources. 3900 Commonwealth

Blvd., Tallahassee, FL 32303
B1LGER, Michael D., 42 Walnut St., Shrewsbury, MA 01545
BILLINGTON, Mark Alan, Box 1327, Friday Harbor, WA 98250
BIRD, Dennis J., 100 Florida St., Apt. 10, Boston, MA 02124
BLACKWELL, Alex H. McCormick, Mamammam Marine Farm Ltd.,

Ross House, Newport, County Mayo, Ireland
BLAKE, Dr. John, 23 Cross Ridge Rd., Chappaqua, NY 10514
BLAKE, Dr. Norman J., Univ. of South Florida, 140 7th Ave.,

St. Petersburg, FL 33701
BLANCHARD, Jean-Andre, Ministre des Peches, P.O. Box 488,

Caraquet, New Brunswick, Canada E0B 1K0
BLOGOSLAWSKI, Dr. Walter, National Marine Fisheries Service,

Northeast Fisheries Center, Milford Lab., Milford, CT 06460
BLUNDON, Jay A., Dept. of Zoology, Univ. of Maryland, College

Park, MD 20742
BOBO, Mildred Yvonne, South Carolina Marine Resources Research

Institute, P.O. Box 12559, Charleston, SC 29412
BOGHEN, Dr. Andrew, Dept. of Biology, Universite de Moncton,

Moncton, New Brunswick, Canada F 1 A 3E9
BONDI, Dr. Kenneth. 14 Jordon Cove Circle, Waterford, CT 06385
BORRERO, Francisco J., Dept. of Biology, Univ. of South Carolina,

Columbia, SC 29208
BOTTON, Mark L., EXCEL Div., Fordham Univ., Collegeof Lincoln

Center, New York, NY 10023
BOUCHET, Phillippe, Museum National d'Histoire Natuelle

Malacologie, 55, Rue de Buffon, 757005 Paris, France
BOURNE, Dr. Neil, Pacific Biological Station, P.O. Box 100.

Nanaimo. British Columbia. Canada V9R 5K6
BRAILSFORD, Paul, Brailsford Associates, 2 Central Street,

Ipswich, MA 01938
BREBER, Paulo, COSPAV, CP. 101, 30015 Chioggia (Venezia) Italia
BREESE, Prof. Wilbur P., Marine Science Center, Marine Science Dr.,

Newport, OR 97365
BRICELJ, V. Monica. Marine Science Research Center, South

Campus Bldg. 6, SUNY-Stony Brook, Stony Brook, NY 11794
BRIGHT, Thomas J., Dept. of Oceanography. Texas A&M Univ.,

College Station, TX 77843
BRITTON, Dr. Joe C, Dept. of Biology, Texas Christian Univ.,

Fort Worth, TX 76129
BROUSSEAU, Dr. Diane J.. Dept. of Biology, Fairfield Univ.,

Fairfield, CT 06430
BROWN, Dr. Carolyn, National Marine Fisheries Service, Milford

Lab, Milford, CT 06460
BROWN, Bradford E.. National Marine Fisheries Service, Southeast

Fisheries Center, 75 Virginia Beach Dr.. Miami, FL 33149
BROWN, Jim, Dept. of Biological Sciences, Simon Fraser Univ.,

Burnaby, British Columbia, Canada V5A 1S6
BUCKNER, Stuart C, Town of Islip, Environmental Management

Div., 577 Main Street, Islip, NY 11751
BUMGARNER, Richard H., Pt. Whitney Shellfish Lab., 1000 Pt.

Whitney Rd., Brinnon, WA 98320
BURCHELL, Edward V., Internet, Inc., 2730 Nevada Ave. N.,

Minneapolis, MN 55427
BUROKER, Dr. Norman E., Oregon Health Sciences Univ. - Dept.

of Biochemistry, 3181 SW Jackson Park Rd.. Portland,

OR 97201

207

208

Membership List - National Shellfisheries association

BURRELL, Dr. Victor G., South Carolina Marine Resources

Research Institute, P.O. Box 12559, Charleston, SC 29412
*BUTLER, Dr. Philip, 106 Matamoros Dr., Gulf Breeze, FL 32561

CAKE, Dr. Edwin W., Jr., Head, Oyster Biology Section, Gulf

Coast Research Laboratory, East Beach Dr., Ocean Springs,

MS 39564
CALABRESE, Dr. Anthony, National Marine Fisheries Service,

Milford Lab, Milford, CT 06460
CAMPBELL, Alan, Biological Station, St. Andrews, New Brunswick,

Canada EOG 2X0
CANZONIER, Walter, 44 Cowart Ave., Manasquan, NJ 08736
CARPENTER, Kirby A., Potomac River Fisheries Commission,

P.O. Box 9, Colonial Beach, VA 22443
*CARRIKER, Dr. Melbourne R.. College of Marine Studies, Univ.

of Delaware, Lewes, DE 19958
CARROLL, William, 509 Bay Dr., Stevensville, MD 21666
CARTER, John A., Martec Ltd., 5670 Spring Garden Rd., Halifax,

Nova Scotia, Canada B3J 1H6
CASTAGNA, Michael, Virginia Institute of Marine Science,

Wachapreague, VA 23480
CASTELL, Dr. John, Department of Fisheries and Oceans, Halifax

Lab, P.O. Box 550, Halifax, Nova Scotia, Canada B3J 2S7
CASTILLO, Silvana, 3 Ave. 12-76. Zona 14, Guatemala City.

Guatemala
CHAISSON, David R., 73 Merrimac Rd., Dartmouth, Nova Scotia,

Canada B2W4W7
CHANLEY, Paul E., P.O. Box 12. Grant, FL 32949
CHATRY, Mark F., Louisiana Dept. of Wildlife & Fisheries, P.O.

Box 37, Grand Isle, LA 70358
CHEN, Ms. Tzyy-Ing, Tungkang Marine Laboratory, Tungkang,

Pingtung, Taiwan 916, Republic of China
*CHESTNUT, Dr. A. F., Institute of Marine Science, Univ. of North

Carolina, Morehead City, NC 28557
CHESTNUT, Dr. A. P.. Biology Dept., Belhaven College. 1500

Peachtree St., Jackson, MS 39202
CHEW, Dr. Kenneth, Div. of Aquaculture and Invertebrate Fisheries,

School of Fisheries, Univ. of Washington, Seattle, WA 98195
CHU, Fu Lin E., Virginia Institute of Marine Science, Gloucester

Point, VA 23062
CLARK, Stephen H., National Marine Fisheries Service, Northeast

Fisheries Center, Woods Hole. MA 02543
CLAYTON, W. E. Lome, Marine Resources Branch, Ministry of

Environment, Parliament Buildings, Victoria, British Columbia.

Canada V8V 1X5
COFFEY, Thomas J., Edgerton Research Lab, New England

Aquarium, Central Wharf, Boston, MA 02110
COLBY, Jean P., 73 Eagle's Nest Rd., Duxbury, MA 02332
COLE, Dr. Timothy J., Horn Point Environmental Lab., Univ. of

Maryland, P.O. Box 775, Cambridge, MD 21613
COLWELL, Dr. R R.. Microbiology Dept., Univ. of Maryland,

College Park, MD 20742
COMMITO, Dr. John, Dept. of Biology, Hood College, Frederick,

MD 21701
CONTE, Dr. Fred S., Aquaculture Extension, Univ. of California.

Davis, CA 95616
CONYERS, James C, Environmental Affairs Group, Potomac

Electric Power Co.. 1900 Pennsylvania Ave. N.W., Washington,

DC 20068
COON, Steven L., Dept. of Zoology, Univ. of Maryland, College

Park, MD 20742
COOPER, Dr. Keith R., School of Pharmacology/Toxicology,

Rutgers Univ., Piscataway, NJ 08854
CORMIER, Paul. 690 Blvd. St. Pierre Quest, Caraquet, New Bruns-
wick, Canada E0B 1K0

COSTA-PIERCE, Barry A., Dept. of Oceanography, Univ. of Hawaii,

Honolulu, HI 96822
COSTLOW, Dr. John D., Duke Univ. Marine Lab. Beaufort, NC 285 16
COVICH, Alan P.. Zoology Dept., 202 Sutton Hall, Univ. of

Oklahoma, Norman, OK 73019
COX, Keith W., 309 Hillside Dr., Woodside, CA 94062
COX, Robert K.,450 KynastonRd.. RR 3, Victoria, British Columbia.

Canada V8X 3X1
CRAIG, Allison, Dept. of Oceanography, Texas A&M Univ., College

Station, TX 77843
CRANCE, Johnie H., U.S. Fish & Wildlife Service, 2625 Redwing

Rd., Ft. Collins, CO 80526
CRAWFORD, Maurice, P.O. Box 286, Woods Hole, MA 02543
CREEKMAN, Laura L., P.O. Box 567, Ilwaco, WA 98624
CRESWELL, R. LeRoy, Center for Marine Biotechnology, Harbor

Branch Institution, Ft Pierce, FL 33450
*CRISP, Dr. Dennis, University College, North Wales, Menai Bridge,

Anglesey, UK
CROCKETT, Lee R., Marine Sciences Institute, Univ. of Connecticut,

Groton, CT 06340
CROSBY, Michael P., Univ. of Maryland, Horn Point Laboratories,

P.O. Box 775, Cambridge, MD 21613
CROWE, Arthur L., Texas Parks & Wildlife Dept., 204 Travis,

Port Lavaca, TX 77979
CUDD, Sue, 2809 165th Place NE, BeUevue, WA 98008
CUMMINS, Joseph M., 4701 W. Maple Lane Circle NW, Gig Harbor,

WA 98335
CUOMO, M. Carmela, Marine Sciences Research Center, State Univ.

of New York, Stony Brook, NY 11794
CUPKA, David M., South Carolina Marine Resource Institute,

P.O. Box 12559, Charleston, SC 29412

DAME, Dr. Richard, Univ. of South Carolina -Coastal Carolina

College, P.O. Box 1954, Conway, SC 29526
DA VIES, Dennis R., ITT Rayonier, Inc.. P.O. Box 299, Hoquiam,

WA 98550
DAVIS, Harold A., Rte. 1. Princess Anne, MD 21853
DAVIS, John D., P.O. Box 156, 25 Old Homestead Rd., Westford,

MA 01886
DAVIS, Jonathan, School of Fisheries VVH-10, Univ. of Washington,

Seattle, WA 98195
DAVIS, Megan, 7600 S.W. 87th Ave., Miami, FL 33173
DAVY, Dr. F. Brian, International Develop. Research Center,

Tjanglin. P.O. Box 101, Singapore 9124
DA WE, Earl G., Dept. of Fish & Oceans, NWAFC, P.O. Box 5667,

St. John's, Newfoundland, Canada A1C 5X1
DAY, Elizabeth Anne, 109-C Thornwell Court, Columbia, SC 29205
DEAN, Dr. David, Box 28, Clarks Cove Rd., Walpole, ME 04573
DeFREESE, Duane E., 933 Waialae Circle NE, Palm Bay, FL 32905
DEMORY, Darrell, Oregon Dept. of Fish and Wildlife, Marine

Science Dr., Newport, OR 97365
deQUILLFELDT, Charles, Marine Sciences Research Center, State

Univ. of New York, Stony Brook, NY 11794
DeVOE, M. Richard, South Carolina Sea Grant Consortium, 221

Fort Jackson Rd., Charleston, SC 29412
DEY, Noel Dean, College of Marine Studies. Univ. of Delaware,

Lewes, DE 19958
DiCOSIMO, Jane, Virginia Institute of Marine Science, Gloucester

Point, VA 23062
DINNEL, Dr. Paul A., Univ. of Washington, Fisheries Research

Institute WH-10, Seattle, WA 98195
DiSALVO, Louis H., Casilla 480, Coquimbo, Chile
DONALDSON, James D., P.O. Box 583, Quilcene, WA 98376
DOWGERT, Martin P., U.S. Food & Drug Admin., 585 Commercial

St., Boon, MA 02108

Membership List - National Shellfisheries association

209

DOWN, Dr. Russel J., Oysterrific. P.O. Box 156, Cape May Court

House, NJ 08210
DOWNING, Sandra L., 1635 33rd Ave., Seattle, WA 98122
DRAZBA, Lawrence, 405 N. Lincoln, Orange. CA 92666
DREDGE, M.. Fisheries Laboratory. Burnett Heads, 4670,

Queensland, Australia
DRESSEL, David, NOAA. National Marine Fisheries Service,

3300 Whitehaven St., NW, Washington. DC 20235
DRINKWAARD, Dr. A. C, Molluscan Shellfish Department,

Julianastraat 18. P.O. Box 135, 1790 AC DenBurg-Texel. The

Netherlands
DRUCKER, Denson, 11667 Newbridge Ct„ Reston, VA 22091
DRURY, Paul E.. 8527 Jennifer No. 5. Juneau, AK 99801
DUBE, Paul.. Marine Sciences Research Center, State Univ. of

New York, Stony Brook, NY 11794
DUGAS, Charles N., 662 E. Perrault St.. Opelousas, LA 705 70
DUGAS, Ronald J.. St. Amant Marine Lab.. Louisiana Dept. of

Wildlife & Fisheries. P.O. Box 37. Grand Isle, LA 70358
DUKE, Dr. Thomas W., U.S. Environmental Protection Agency

Lab., Sabine Island. Gulf Breeze. FL 32561
DUNCAN, Dr. Patricia. College of William & Mary. Virginia Institute

of Marine Science, Gloucester Point, VA 23062
DUNNINGTON, Elgin, Chesapeake Biological Lab., Box 523,

Solomons, MD 20688
DURFEE, Dr. Wayne K„ 44 Bridgetown Rd„ Sunderston, RI 02874

EATON, Jonathan F., 4- A Gleason St., Thomaston, ME 04861
EBERT, Earl E.. California Dept. of Fish and Game, Granite Canyon

Coast Route, Monterey, CA 93940
EBLE, Dr. Albert F., R.D. No. 6. Box 345-B, Flemington, NJ 08822
ECKMAYER, William J., Alabama Dept. of Conservation and

Natural Resources, Marine Resources Div., P.O. Box 189,

Dauphin Island, AL 365 28
EDWARDS, Dr. D. Craig, Univ. of Massachusetts, Zoology Dept.,

Amherst, MA 01003
EDWARDS, Sarah B., Pine Lane, Barstable, MA 02630
EINOLF, David M., 1817 W. Call St., Apt. F-8, Tallahassee, FL 23204
EISELE, William J., New Jersey Div. of Water Resources, Leeds

Point Field Office, Star Rte., Abescon, NJ 08201
EISLER, Dr. Ronald, U.S. Fish & Wildlife Service, Patuxent Wildlife

Research Center, Laurel, MD 20708
ELDRIDGE, Peter J., 761 Stiles Dr., Charleston. SC 29412
ELLIFRIT, N. J., 16217 NE 22nd Ave., Ridgefield, WA 98642
ELLIOT, Elisa L., Dept. of Microbiology, Univ. of Maryland,

College Park, MD 20742
ELLIS, Dr. Derek, Biology Dept., Univ. of Victoria, Victoria,

British Columbia, Canada V8W 2Y2
ELNER, Dr. Robert W., Fisheries & Oceans, Biological Station.

St. Andrews, New Brunswick, Canada E0G 2X0
ELSKUS, Adria A.. School of Oceanography, Univ. of Rhode Island,

Kingston, RI 02881
ELSTON, Dr. Ralph, Battelle Marine Research Lab.. 439 Sequim

Rav Rd.. Sequim, WA 98382
EMERY, Ann, 3421 Shepherd St., Chevy Chase, MD 20815
ENRIGHT, Dr. Catherine. Ketch Harbour, Halifax County, Nova

Scotia, Canada B0J 1X0
EPP, Jennifer. Marine Sciences Research Center, State Univ. of

New York, Stony Brook, New York 11794
ERICKSON, Jeffery T.. Univ. of Miami, Rosenstiel School of

Marine and Atmospheric Science, Div. of Biological and Living

Resources, 4600 Rickenbacker Causeway, Miami. FL 33149
EVANS, Camille, P.O. Box 731. Quilcene, WA 98376
EVERSOLE, Dr. Arnold B., Dept. of Aquaculture. Fisheries &

Wildlife, 310 Long Hall, Clemson Univ., Clemson, SC 29631
EWALD, Joseph Jay. Apartado 1198, Maracaibo, Venezuela

FEDER, Dr. Howard, Institute of Marine Science. Univ. of Alaska,

Fairbanks, AK 99701
FENG, Dr. SungY.. Marine Sciences Institute, Univ. of Connecticut,

Groton, CT 06340
FERGUSON, Ernest, P.O. Box 488, Caraquet, New Brunswick,

Canada E0B 1K0
FERNANDEZ, Gustavo E., College of Marine Studies, Univ. of

Delaware, 700 Pilottown Rd., Lewes, DE 19958
FISHER, William S., Univ. of Maryland, Horn Point Laboratories,

P.O. Box 775, Cambridge, MD 21613
FITZGERALD, Lisa M.. Univ. of Miami, Rosenstiel School of

Marine and Atmospheric Science. Div. of Biological and Living

Resources, 4600 Rickenbacker Causeway, Miami. FL 33149
FLAGG, Paul J., 31 Kings Point Rd., East Hampton, NY 11937
FLICK, Dr. George J.. Food Science & Technology Dept., Virginia

Polytechnic Inst.. Blacksburg. VA 24061
♦FLOWER, H. Butler, F. M. Flower & Sons, P.O. Box 1436, Bayville,

NY 11709
FOLLET, Jill E.. Alaska Dept. of Fish and Game, 333 Raspberry

Rd., Anchorage, AK 99502
FOLTZ, David W., Dept. of Zoology & Physiology, Louisiana State

Univ., Baton Rouge, LA 70803
FORBES, Dr. Milton, College of the Virgin Islands, P.O. Box 206,

Kingshill, St. Croix, VI 00850
FORD, Dr. Susan E., Rutgers Univ. Center Research Lab., Box 587,

Port Norris, NJ 08349
FOSTER, Walter S., P.O. Box 637. Hatchet Cove. Friendship, ME

04547
FOX, Richard, New York Dept. of Environmental Conservation,

Bldg. 40, State Univ. of New York, Stony Brook, NY 11794
FREEMAN, Dr. John A., Dept. of Biology. Univ. of South Alabama,

Mobile, AL 36688
FRITZ, Lowell W., Rutgers Univ. Oyster Research Lab., P.O. Box

587, Port Norris, NJ 08349
FRULAND, Robert M., 7128 South Shore Dr., South Pasadena, FL

33707
FULLER, Sue Cynthia, Dept. of Zoology, Rutgers Univ., Box 1059,

Piscataway, NJ 08854
FYFE, David A., 155-7072 Inlet Dr.. Burnaby. British Columbia.

Canada V5A 1C2

GAFFNEY, Patrick M., Dept. of Ecology and Evolution, State

Univ. of New York, Stony Brook, NY 1 1 794
GAILEY, Matthew D., Juniper Point Sea Farms, 3 Juniper Point Rd.,

Branford, CT 06405
GALLAGER, Scott M., Woods Hole Oceanographic Institution,

Woods Hole, MA 02543
GALLANT, W. E., Snow Food Products, P.O. Box F. Old Orchard

Beach, ME 04064
GANGMARK, Carolyn E., P.O. Box 549, Manchester, WA 98353
GAREY, John F., 65 Olde Knoll Rd., Marion. MA 02738
CARLO, Elizabeth V, Battelle Research Laboratory, P.O. Drawer

AH. Dux bury. MA 02332
GARREIS, Mary Jo, 129 Severn Way, Arnold, MD 21012
GATES, Keith W.. Univ. of Georgia Marine Extension Service,

P.O. Box Z, Brunswick. GA 31521
GEOGHEGAN, Paul, 28 Williams St.. Salem, MA 01970
GEORGE, Keith, Agridex Ltd.. 47 Mowbray Rd., Northallerton.

North Yorkshire. England DL6 1QT
GERRIOR, Patricia. National Marine Fisheries Service, Emerson

Ave., Gloucester, MA 01930
GIBBONS, Dr. Mary C, College of William & Mary, Virginia Institute

of Marine Science. Wachapreague, VA 23480
GIBSON, Dr. Charles I., Battelle Memorial Institute, 505 King Ave.,

Columbus, OH 43201

210

Membership List - National Shellfisheries Association

GLENN, Dr. Richard D., 1704 Gotham St., Chula Vista, CA 92010
*GLUDE, John B., 2703 W. McGraw St., Seattle, WA 98199
GOLDBERG, Ronald, National Marine Fisheries Service. Milford

Lab., Milford, CT 06460
GOOD, Lorna, 128 Hitchner Hall, Univ. of Maine, Orono, ME 04469
GOODGER, Timothy E., National Marine Fisheries Service, Oxford

Lab., Oxford, MD 21654
GOODSELL, Joy G., Rutgers Univ., Shellfish Research Lab., Box

587, Port Norris, NJ 08349
GOODWIN, Lynn, Pt. Whitney Shellfish Lab., 1000 Pt. Whitney Rd..

Brinnon, WA 98320
GOULD, Edith, National Marine Fisheries Service, Milford Lab.,

212 Rogers Ave., Milford, CT 06460
GRAY, C. Scott, 411 Liberty St., Santa Cruz, CA 95060
GREEN, William C, 64 Leetes Island Rd., Guilford, CT 06437
GREENE, Gregory T., 123 Bay Ave., Bayport, NY 11705
GRIM, John S., Northeastern Biological, Inc., Kerr Rd., RD 3,

Rhinebeck, NY 12572
GRISCHKOWSKY, Dr. Roger S., Alaska Dept. of Fish and Game,

333 Raspberry Rd., Anchorage, AK 99502
GRUBER, Gregory L„ Dept. of Health & Hygiene, Office of

Environ. Programs, 415 Chinquapin Round Rd., Annapolis, MD

21401
GRUBLE, Edward J., 8622 Fauntlee Crest SW, Seattle, WA 98136
*GUNTER, Dr. Gordon, Director Emeritus, Gulf Coast Research Lab.,

Ocean Springs, MS 39564
GUSSMAN, David S., Virginia Institute of Marine Science, College

of William & Mary, Gloucester Point, VA 23062

HADLEY, Nancy H., 1214 Grimsley Dr., Charleston, SC 29412
HALLDORSON, Dori. Coast Oyster Co., Box 166, South Bend,

WA 98586
HAMM, Gerald L., 10563 NW 2nd Court, Plantation, FL 33324
HAMMERSCHMIDT, Paul C, 1821 Algee, Port Lavaca, TX 77979
HAMMERSTROM, Richard J., 2901 Shamerock South, Tallahassee,

FL 32308
HANKS, Dr. James E., P.O. Box 253, Milford, CT 06460
HARGIS, Dr. William J., Jr., Virginia Institute of Marine Science,

College of William and Mary, Gloucester Point, VA 23062
HARRIS, Robert E., Virginia Institute of Marine Science-Jefferson

Hall, College of William and Mary, Gloucester Point, VA 23062
HARTSELL, James A., 15 Chester St., Apt. 1, New London, CT

06320
HARTWICK, Dr. Brian, Dept. of Biological Science, Simon Fraser

Univ., Burnaby, British Columbia, Canada V5A 1S6
HASELTINE, Arthur W., Marine Culture Lab.. Granite Canyon,

Coast Route, Monterey, CA 93940
*HASKIN, Dr. Harold H., Dept. of Oyster Culture, Rutgers Univ.,

P.O. Box 1059, Piscataway, NJ 08854
HAVEN, Dexter S., Virginia Institute of Marine Science, College of

William and Mary, Gloucester Point, VA 23062
HAXBY, Richard E., c/o Morton Bahamas Ltd., Matthewtown,

Inagua, Bahamas
HAYDEN, Barbara J., Fisheries Research Div., P.O. Box 297,

Wellington. New Zealand
HEARD, Dr. Richard, P.O. Box 878. Ocean Springs, MS 39564
HEIDEMAN, Robert. P.O. Box 1446, Apopka, FL 32704
HEINEN, Dr. John M., Dept. of Wildlife and Fisheries, P.O. Drawer

LW, Mississippi State, MS 39762
HELM, Nancy E., Marine Sciences Research Center, State Univ. of

New York, Stony Brook, NY 1 1794
HENDERSON, Bruce Alan, Marine Sciences Center, Oregon State

Univ., Newport, OR 97365
HENSEN, Roberto, Fonds Caracopreoject, P.O. Box 43, Bonaire,

Netherlands Antilles

HEPWORTH, Daniel A., Rt. 3, Box 135, Hayes, VA 23072
HERITAGE, G. Dwight, Pacific Biological Station, Nanaimo,

British Columbia, Canada V9R 5K6
HERRMANN, Robert B., 101 King St., New Bern, NC 28560
HERSHBERGER, Dr. William K., School of Fisheries WH-10.

Univ. of Washington, Seattle, WA 98195
HICKEY, John M., Massachusetts Div. of Marine Fisheries, 449

Route 6Ah, East Sandwich, MA 02537
HICKEY, Mary T., 4415 Independence St., Rockville, MD 20853
HIDU, Dr. Herbert, Ira C. Darling Center, Univ. of Maine, Walpole,

ME 04573
HILLMAN, Dr. Robert E., Battelle New England Marine Research

Laboratory, Washington St., Duxbury, MA 02332
HIRSCHBERGER, Wendy, 5832 NE 75th. No. 205. Seattle. WA

98115
HOCHHEIMER, John N., Marine Advisory Program, Univ. of

Maryland, CEES, P.O. Box 775, Cambridge, MD 21613
HOENIG, John M., Minnesota Dept. of Natural Resources, Box 25,

Centennial Office Bldg., St. Paul, MN 55 155
HOESE, Dr. H. Dickson, Dept. of Biology. Univ. of Southwestern

Louisiana, Lafayette, LA 70501
HOFSTETTER, Robert P.. Rt. 1,4831 Elm St., Seabrook, TX 77586
HOLMES, Patrick B., P.O. Box 2651, Kodiak, AK 99615
HOOPER, Craig, 214 Meadowlook Way, Boulder, CO 80302
*HOPKINS, Dr. Sewell H., Biology Dept., Texas A&M Univ.. College

Station, TX 77843
HOPKINS, Steve, WaddeU Mariculture Center, P.O. Box 809,

Bluffton. SC 29910
HORTON, Dr. Howard F., Fisheries & Wildlife Dept., Oregon

State Univ., Corvallis, OR 97331
HOUGHTON, Jonathan, Dames and Moore, 155 NE 100th, Seattle,

WA 98125
HOUK, James L., California Dept. of Fish and Game, Marine Culture

Lab., Granite Canyon Coast Route, Monterey, CA 93940
HOWELL, Robert. Dept. of Biology, Conradi Bldg.. Florida State

Univ., Tallahassee, FL 32306
HOWSE, Dr. Harold D., Gulf Coast Research Laboratory, Ocean

Springs, MS 39564
HRUBY, Thomas, RCA, 159 Main St., Gloucester, MA 01930
HRUSE, Michael W., RD 1. Box 165, Fire Lane, Vincentown, NJ

08088
HUBER, L. Albertson, Back Neck Rd., Rte. 4, Bridgeton, NJ 08302
HUGUENIN, John E., 49 Oyster Pond Rd., Falmouth, MA 02540
HUMPHREY, Celeste, Dalton Junior College, Dalton, GA 30720
HUNER, Dr. Jay V., 1144 Rue Crozat, Baton Rouge, LA 70810
HUTCHISON, F. M., P.O. Box 281, Cayucos, CA 93430

IBARRA, Ana Maria, Dept. of Fisheries and Wildlife. Oregon State

Univ., Corvallis, OR 97331
INCZE, Dr. Lewis S., NWAFC/RACE Div., 7600 Sand Pt. Way, NE,

BIN C15700, Seattle, WA 981 12
INGLE, Donna M., Rt. 16, Box 9034, Tallahassee, FL 32304
INGLE, Robert M. 173 Avenue B, Apalachicola, FL 32320
IVERSEN, Dr. Edwin S., Univ. of Miami, Rosenstiel School of

Marine and Atmospheric Science, Div. of Biological and Living

Resources, 4600 Rickenbacker Causeway, Miami. FL 33149

JEFFERDS, Peter, Penn Cove Mussels, Inc., P.O. Box 148, Coupe-

ville, WA 98239
JENNINGS, Charles R.. P.O. Box 5620, Berkeley, CA 94705
JEWELL, Dr. Sheila Stiles, National Marine Fisheries Service,

Milford Lab., 212 Rogers Ave., Milford, CT 06460
JEWETT, Stephen, Institute of Marine Science. Univ. of Alaska,

Fairbanks, AK 99701
JOHNSON, Scott, 5736 CessnaAve., Apt. W,FridayHarbor,WA98250

Membership List - National Shellfisheries association

211

JONES, Gordon B., Skerry Bay, Lasqueti Island, British Columbia,
Canada VOR 2J0

JONES, Dr. Douglas S., Dept. of Geology, Univ. of Florida, Gaines-
ville, FL 32611

JORY, Darryl E., Univ. of Mainii, Rosenstiel School of Marine and
Atmospheric Science, Div. of Biological and Living Resources,
4600 Rickenbacker Causeway, Miami. FL 33149

JOYCE, Edwin A.. Jr.. 14130 N. Meridian Rd., Tallahassee, FL 32312

JUDSON, Irwin, P.O. Box 2000, Charlottetown, Prince Edward
Island, Canada CIA 7N8

KAMENS, Todd C, College of Marine Studies, Univ. of Delaware,

700 Pilottown Rd., Lewes. DE 19958
KANE, Dr. Bernard, Dept. of Environmental Health, East Carolina

Univ., Greenville, NC 27834
KARINEN, John F„ Auke Bay Biological Lab., P.O. Box 210155,

Auke Bay, AK 99821
KARNEY, Richard C, Box 1552, Oak Bluffs, MA 02557
KASSNER, Jeffrey, 28 Penn Commons, Shiiley, NY 11967
KEAN, Joan, Fisheries and Oceans. 1707 Lower Water St.. Halifax,

Nova Scotia, Canada B3J 2S7
KEITH, W. J., South Carolina Marine Resources Research Institute,

P.O. Box 12559, Charleston, SC 29412
KELLER, Thomas E., Box 285, RR No. 1, Edgecomb, ME 04556
KELP1N, Geraldine, 329 East State St., Long Beach. NY 11561
KENNEDY, Dr. Victor S., Horn Point Environmental Lab., Box 775,

Cambridge, MD 21613
KENNISH, Dr. Michael J., GPU Nuclear, Oyster Creek Nuclear

Station, P.O. Box 388, Forked River, NJ 08731
KENSLER, Dr. Craig B., UNESCO Marine Science Project (UNDP

POUCH, Rangoon, Burma), UNDP/One United Nations Plaza,

New York, NY 10017
KILGEN, Marilyn B., Dept. of Biological Sciences, Nicholls State

Univ., Thibodaux, LA 70310
KILGEN, Dr. Ronald H., Dept. of Biological Sciences, Nicholls

State Univ., Thibodaux, LA 70310
KLINE, Thomas C, School of Fisheries WH-10, Shellfish Unit,

Univ. of Washington, Seattle, WA 98195
KNAUB, Richard S., Dept. of Aquaculture, Fisheries and Wildlife,

Clemson Univ., Clemson, SC 29631
KOGANEZAWA,Akimitsu, Aquaculture Div., Tohoku Reg. Research

Lab., 3-27-5, Shinhamacho, Shiogama, Miyagi-Ken 985 Japan
KOPPELMAN, Lee E., Long Island Regional Planning Board,

Veterans Memorial Highway, Happauge, NY 11788
KRAEUTER, Dr. John N., Baltimore Gas & Electric Co., Crane

Aquaculture, P.O. Box 1475, Baltimore, MD 21203
KRANTZ, David E., Marine Science Program, Univ. of South

Carolina, Columbia. SC 29208
KRAUS, Richard A., Aquaculture Research Corp., P.O. Box AC,

Dennis, MA 02638
KRYGSMAN, Adrian, 35 Madeline Ave., Clifton, NJ 0701 1
KUNKLE, Donald E„ Rutgers Univ. Oyster Research Lab., P.O.

Box 587, Port Norris, NJ 08349
KURKOWSK1, Kenneth P., 234 Fenimore Ave., Uniondale. NY

11553
KUTRUBES, Leo P., National Labs, 114 Waltham St., Lexington,

MA 02173
KYTE, Michael A., 527 212th St., SW, Bothell. WA 98021

LANDRUM, Michael R., 902 S.E. Belfast Ave., Port St. Lucie,

FL 33452
LANGDON, Dr. Chris, College of Marine Studies. Univ. of Delaware.

Lewes, DE 19958
LANGE, Anne M. T., National Marine Fisheries Service, Northeast

Fisheries Center, Woods Hole, MA 02543

LANGTON, Richard W., Marine Research Lab., Dept. of Marine

Resources, West Boothbay Harbor, ME 04575
LAVOIE, Dr. Rene E., Dept. of Environment, Fisheries Service,

P.O. Box 550, Halifax, Nova Scotia, Canada B3J 2R3
LAWDER, HanyC.,512 8th Street, Port St. Joe, FL 32456
LAWTON, Peter, Dept. of Fisheries & Oceans, Biological Station,

St. Andrews, New Brunswick, Canada E0G 2X0
LEARY, Terrance R., Gulf of Mexico Fisheries Management Council,

5401 W. Kennedy, Suite 881, Tampa, FL 33609
LEIB, Susanne, Florida Institute of Technology, Box 339, Jensen

Beach, FL 3345 7
LE1BOVITZ, Dr. Louis, Director, Laboratory for Marine Animal

Health, Marine Biological Laboratory. Woods Hole, MA 02543
LESLIE, Mark D., 5 Deborah St.. Waterford, CT 06385
s LINDSAY, Cedric E., 560 Pt. Whitney Rd.. Brinnon, WA 98320
LINDSAY, John A., P.O. Box JJ, Durham, NH 03824
LIPOVSKY, Vance P., P.O. Box 635, Ocean Park, WA 98640
LITTLE, Edward J., Florida Dept. of Natural Resources, P.O. Box

404, Key West, FL 33040
LIVINGSTON, Dr. Robert J., Dept. of Biological Science, Florida

State Univ., Tallahassee, FL 32306
LOCKWOOD, George S., Monterey Abalone Farms, 300 Cannery

Row, Monterey, CA 93940
LOGUE, Maureen D., Ira C. Darling Center, Univ. of Maine,

Walpole, ME 04573
LOMAX, Dr. Ken, Dept. of Agricultural Engineering. Univ. of

Delaware, Newark, DE 19711
*LOOSANOFF, Dr. Victor L., 17 Los Cerros Dr., Greenbrae, CA

94904
LORING, Richard H.. Aquacultural Research Corp., P.O. Box AC,

Dennis, MA 02638
LOUGH, Dr. Robert G., National Marine Fisheries Service, North-
east Fisheries Center, Woods Hole, MA 02543
LOVELAND, Robert E., Dept. of Zoology, Rutgers Univ., P.O. Box

1059, Piscataway, NJ 08854
LOWE, Jack I„ Route 2, Box 20, Gulf Freeze, FL 32561
LUBET, Prof. Pierre, Laboratoire de Zoologie, Universite' de

Caen 14032. Caen Cedex, France
LUTZ, Rebecca Ashley, 52 Main St., P.O. Box 215, Bloomsbury,

NJ 08804
LUTZ, Dr. Richard A., Oyster Research Laboratory, Rutgers

Univ., P.O. Box 1059, Piscataway, NJ 08854

MacDONALD, Dr. Bruce, Pacific Biological Station, P.O. Box 100,

Nanaimo, British Columbia, Canada V9R 5K6
MacFARLANE, Sandra Libby, Orleans Shellfish Dept., Orleans,

MA 02653
MacKENZIE, Clyde L., Sandy Hook Laboratory, Highlands, NJ

07732
MacLEOD, Lincoln-Lowell, P.O. Box 700, Pictou, Nova Scotia,

Canada B0K 1H0
MAGOON, Charles D., Dept. of Natural Resources, Marine Land

Management. Olympia, WA 98504
M ALONE, Ronald F., Dept. of Civil Engineering, Louisiana State

Univ., Baton Rouge, LA 70803
MALOUF, Dr. Robert. Marine Sciences Research Center, State

Univ. of New York, Stony Brook, NY 11794
MANDRUP-POULSEN, Jan, Dept. of Oceanography, Florida State

Univ., Tallahassee, FL 32306
MANN, Dr. Roger, Woods Hole Oceanographic Institution, Woods

Hole, MA 02543
MANZI, Dr. John J., Marine Resources Research Institute, P.O.

Box 12559, Charleston, SC 29412
MARIS, Robert, P.O. Box 6322, Norfolk, VA 23508
MARSHALL, Dr. Nelson, P.O. Box 1056, St. Michaels, MD 21663

212

Membership list - National Shellfisheries association

MARSHALL, Howard L., Environmental Protection Agency, 345

Courtland St., NE, Atlanta, GA 30365
MARSTON, Claudie L., 33 Nichols Ave., Apt. 3, Newmarket, NH

03857
MARTIN, Roy E., National Fisheries Institute, 2000 M St., NW,

Suite 580, Washington, DC 20036
MARU, Dr. Kuniyoshi, Hokkaido Institute of Maticulture, Shikabe,

Hokkaido. 041-14, lapan
MASON, Katherine. 217 Murray Hall, Univ. of Maine, Orono, ME

04469
MAUGLE, Paul D., 323 Graduate Village, Kingston. RI 02881
MAYER, Marianne, Marine Extension, Univ. of Georgia, P.O. Box

13687, Savannah, GA 31406
McBETH, Dr. James W., P.O. Box 1540, Carlsbad, CA 92008
MCCARTHY, Charles, 2000 Cox Neck Rd., Mattituck, NY 11952
McCONAUGHA, Dr. John R., Dept. of Oceanography, Old Dominion

Univ., Norfolk, VA 23508
McCUMBY, Kristy I., 2590 Lingonberry Lane, Fairbanks, AK 99701
McEWEN, Laurel A., General Delivery, Nahcotta, WA 98637
McFADDEN, Murray, 577 W. 28th Ave., Vancouver, British

Columbia, Canada V5Z 2H2
McGRAW, Dr. Katherine A, 131 N. 40th, Seattle, WA 98103
McHUGH, Dr. J. L., Marine Sciences Research Center, State Univ.

of New York, Stony Brook, NY 1 1 794
McLAUGHLIN, Dave. Agricultural Engineering Dept., Clem son

Univ., Clemson, SC 29631
McMURRER, Kathleen A.. 36 Woodfield Ave., Fort Salonga, NY

11768
McTEER, Temple, Waddell Mariculture Center, P.O. Box 809,

Bluff ton, SC 29910
*MEDCOF, Dr. J. C, P.O. Box 83, St. Andrews, New Brunswick,

Canada E0G 2X0
*MENZEL, Dr. R. Winston. Dept. of Oceanography, Florida State

Univ., Tallahassee, FL 32306
MERCALDO. Renee S., National Marine Fisheries Service. Rogers

Ave., Milford, CT 06460
MERCER, Dr. J. P.. Shellfish Research Lab., Carna, County Galway.

Ireland
MERRILL, Dr. Arthur S., 25 North Front St., Richmond, ME 04357
MIANMANUS, Ratsuda, Univ. of Miami. Rosenstiel School of

Marine and Atmospheric Science, Div. of Biological and Living

Resources, 4600 Rickenbacker Causeway. Miami. FL 33149
MIDDLETON, Karen Chandler, 175 Abram Hill Rd.. Duxbury, MA

02332
MILLER, George C, 16140 S.W. 108th Court, Miami, FL 33157
MILLER, Mum, Route 1, Bowler, WI 54416
MILLER, Robert E., P.O. Box 775, Cambridge, MD 21613
MILLER, R. J., P.O. Box 550, Halifax, Nova Scotia, Canada B3J 2S7
MILMOE, Gerard F., Box 446, Port Jefferson, NY 11777
MIX, Dr. Michael C, General Science Dept., Weniger Hall 355,

Oregon State Univ., Corvallis. OR 97330
MOORE, M. Mug, Mercenaria Manufacturing, R.D. 1. Box 293-B,

Millsboro, DE 19966
MORADO, J. Frank. RACE Div., Bldg. 4, Rm. 2083. 7600 Sand

Point Way, NE, BIN-C15700, Seattle, WA 98115
MORGAN, Dr. Bruce H., P.O. Box 8811, Portland, OR 97207
MORIYASU, Mikio, Marine Biology Research Center, Univ. of

Moncton, Moncton, New Brunswick, Canada E1A 3E9
MORRISON, Allan, 95 Scott St., Charlottetown. Prince Edward

Island, Canada C1E 1A1
MORRISON, George, Environmental Protection Agency, South

Ferry Rd., Narragansett, RI 02882
MORSE, Dr. M. Patricia, Marine Science Institute, Northeastern

Univ., Nahant, MA 01908
MOSS, Charles G.. Rt. 2 Armory, Angleton, TX 77515

MOSS, Shaun, 570 Pilottown Rd., Lewes, DE 19958

MUISE, Brian, P.O. Box 84, Musquodoboit Harbour, Nova Scotia,

Canada B0J 2L0
MULVIHILL, Michael, AREA, P.O. Box 1303, Homestead, FL 33090
MUNDREN, Fentress, North Carolina Div. of Marine Fisheries,

P.O. Box 769, Morehead City, NC 2855 7
MURPHY, Richard C, The Cousteau Society, 8439 Santa Monica

Blvd., Suite 1 10, Los Angeles, CA 90069
MURPHY, William A.. P.O. Box 1236, Charlottetown, Prince

Edward Island, Canada CIA 7M8
MURRAY, Robert L., 6211 SW 79th St., Miami, FL 33143
MUSGROVE, Nancy A., College of Fisheries, Univ. of Washington,

Seattle, WA 98195

NAKAGAWA, Yoshihiko, Hokkaido Hakodate Fisheries Experiment

Station, Ynokawa-cho 1-cho 2-66. Hakodate, Hokkaido. Japan
NAKAL, Alberto, 4223 SW 6th, Miami, FL 33134
NASSER, Sergio E. Rivera, Apartado Postal 749, Cuidad Obregon,

Sonora, Mexico
NEAL.Dr. Richard, c/o Gilbert Neal, Box 623, Shell Rock, I A 50670
NEIMA, Paul G., Fisheries Resource Dev., Ltd., 192 Joseph Zatzman

Dr. S., 192, Dartmouth, Nova Scotia, Canada B3B 1N4
NELSON, Chris, Marine Sciences Research Center, State Univ. of

New York, Stony Brook, NY 11794
NELSON, David A.. National Marine Fisheries Service, Milford,

CT 06460
"NELSON, J. Richard, 371 Post Rd., Madison, CT 06443
NEUDECKER, Dr. Thomas. Inst, fur Kusten- und Binnenfischerei;

Aussenstelle Langballigau, Am Hafen D-2391 Langballig, West

Germany
NEWBERG, Douglas, College of Marine Studies, Univ. of Delaware,

700 Pilottown Rd., Lewes, DE 19958
NEWELL, Carter R., Maine Shellfish Research and Development,

RFD 1, Box 149, Damariscotta, ME 04543
NEWELL, Dr. Roger 1. E.. Horn Point Environmental Lab., Univ.

of Maryland, P.O. Box 775. Cambridge. MD 21613
NEWKIRK, Dr. Gary F., Biology Dept.. Dalhousie Univ., Halifax,

Nova Scotia, Canada B3H 4J1
NORMAN-BOUDREAU, Karen, Bodega Marine Laboratory, Bodega

Bay, CA 94923
NORRIS, Robert M., Potomac River Fisheries Comm., 222 Taylor

St., Colonial Beach. VA 22443
NOSHO, Terry Y, 12510 Langston Road, South, Seattle, WA 98178
NOVOTNY, Anthony, 1919 E. Calhoun, Seattle, WA 98112

OAKES, Diane R.. Laboratory for Experimental Biology. National
Marine Fisheries Service, 212 Rogers Ave., Milford, CT 06460

O'BRIEN, Dr. Francis X., Dept. of Biology, Southeastern Massa-
chusetts Univ., North Dartmouth, MA 02747

O'BRIEN, Loretta, P.O. Box 597, Woods Hole, MA 02543

O'DOR. Dr. Ronald K., Biology Dept., Dalhousie Univ.. Halifax,
Nova Scotia, Canada B3H 4J1

OESTERLING, Michael J., Virginia Institute of Marine Science,
College of William and Mary, Gloucester Point, VA 23062

OLMI, Eugene J., Grice Marine Biological Lab., Collegeof Charleston,
Charleston.SC 29412

OLSEN, Dr. Lawrence A., Florida Dept. of Environmental Regula-
tion, 2600 Blairstone Rd.. Tallahassee, FL 32301

OLSEN, Scharleen, Washington Dept. of Fisheries, 1000 Pt. Whitney
Rd., Brinnon, WA 98320

OSIS, Laimons, Oregon Fish Comm., Marine Science Dr., Newport,

O OR 97365

OTWELL, Dr. W. Steven, Food Science and Human Nutrition, Univ.
of Florida, Gainesville, FL 326 1 1

Membership List - National Shellfisheries association

213

OVERSTREET, Dr. Robin M., Gulf Coast Research Laboratory,

Ocean Springs, MS 39564
OVS1ANICO, Natalya N.,c/o Morton Bahamas Ltd., P.O. Box 1216,

Brunswick, GA 31521

PAGE, Mark. Marine Science Institute, Univ. of California, Santa

Barbara, C A 93106
PAGEL, Robert, 5 South Grand Ave., Deerfield, WI 53531
PAUL, Augustus John. Seward Marine Station, Institute of Marine

Science, Box 617, Seward, AK 99664
PEARCE, Dr. John B., National Marine Fisheries Service, Sandy

Hook Laboratory, Highlands, NJ 07732
PEIRSON, W. Michael, P.O. Box 222, Eastville,VA 23347
PENNER, Dr. Lawrence R., Biological Science Group U-43, Univ. of

Connecticut, Storrs, CT 06268
PERDUE, James A., 1 709 Upper Millstone Lane, Salisbury , MD 21801
PERLMUTTER, Dr. Alfred, Biology Dept., New York Univ., New

York, NY 10012
PERRY, Harriet M, Gulf Coast Research Laboratory, Ocean Springs,

MS 39564
PETROVITZ, Eugene J, Aquacultural Research Corp., P.O. Box AC,

Dennis, MA 02638
PFITZENMEYER, Hayes T., Chesapeake Biological Laboratory,

Box 38, Solomons, MD 20688
PHILLIPS, Clyde A., High & Rena Streets, Mauricetown, NJ 08329
PILLSBURY, Katherine, A2-27 Twin Oaks Village, Mansfield,

MA 02048
POBRAN, Theodore T. , Marine Resources Branch, 229-780 Blanchard

Street. Victoria. British Columbia, Canada V8V 1X5
POIRRIER, Dr. Michael A., Dept. of Biological Sciences, Univ. of

New Orleans, Lake Front, New Orleans, LA 70148
PONDICK, Jeffrey, Biological Sciences Group, Univ. of Connecticut,

Storrs, CT 06268
POPHAM, Dr. J. David, Seakem Oceanographic, Ltd., 2045 Mills

Road, Sydney, British Columbia, Canada V8L 3S1
PORTER, Hugh J.. Univ. of North Carolina, Institute of Marine

Science, Morehead City, NC 2855 7
POWELL, Eric N., Dept. of Oceanography, Texas A&M Univ.,

College Station, TX 77843
POWELL, Guy C, Fishery Research Biologist, Box 2285, Kodiak,

AK 99615
PRAKASH, Dr. A., Environmental Protection Service. Place Vincent

Massey; 12th Fir., Ottawa, Ontario, Canada K1A 1C8
PREZANT, Dr. Robert S., Dept. of Biology, Univ. of Southern

Mississippi, Southern Station, Box 5018, Hattiesburg, MS 39401
PRICE, Thomas J., National Marine Fisheries Service, Beaufort, NC

28516
PROCHASKA, Dr. Fred J., Food & Resource Economics, 1170

McCarty Hall, Univ. of Florida, Gainesville, FL 32611
PRUDER, Dr. Gary D., College of Marine Studies, Univ. of

Delaware, Lewes, DE 19958

*QUAYLE, Dr. Daniel B., Fisheries and Oceans, Pacific Biological
Station, Nanaimo, British Columbia, Canada V9R 5K6
QUIN, Judith. 1567 Whiffen Spit. Sooke, British Columbia, Canada

RAE, Dr. John G., Dept. of Natural Science. Florida Institute of

Technology, Jensen Beach, FL 33457
RANEY, Dr. Edward C, 301 Forest Dr., Ithaca, NY 14850
RASK, Hauke, P.O. Box 209, Barnstable, MA 02630
RATHJEN, Warren F., P.O. Box 1109, Gloucester, MA 01930
RAYLE, Michael F., Steimle & Associates, Inc., P.O. Box 856,

Metairie, LA 70004
REISINGER, Tony, Cameron County Extension Service, County

Bldg., San Benito, TX 78586
RELYEA, David R., F. M. Flower & Sons, Inc., 34 Ludlum Ave.,

BayviUe, NY 11709
RHODES, Bryce W., 3190 A Airport Loop Dr., Costa Mesa, CA

92626
RHODES, Dr. Edwin, National Marine Fisheries Service, 212 Rogers

Ave., Milford, CT 06460
RHODES, Raymond, 8 Westside Dr., Charleston, SC 29412
RICE, Mindy L., 43 Larkin St., Bangor. ME 04401
RIDEOUT, Carol B., Virginia Institute of Marine Science, College

of William and Mary, Gloucester Point, VA 23062
RINES, Henry M.. School of Oceanography, Univ. of Rhode Island,

Kingston, RI 02881
RING, Gregg, P.O. Box 13396, Houston, TX 77219
RIVARA, Gregg, 41 Amagansett Dr., Sound Beach, NY 11789
ROACH, David A., Westport Shellfisheries, Town Hall, 816 Main

St.,Westport. MA02790
ROBERT, Ginette, Fisheries Research Branch, P.O. Box 550.

Halifax, Nova Scotia. Canada B3J 2S7
ROBERTS, Dr. Morris H.. Virginia Institute of Marine Science.

College of William and Mary, Gloucester Point, VA 23062
ROBERTSON, Robert, Dept. of Malacology, Academy of Natural

Sciences. Nineteenth & the Parkway, Philadelphia, PA 19103
ROBINSON, Anja. P.O. Box 312, Y achats, OR 97498
ROBINSON, Dr. William E., New England Aquarium, Edgerton

Research Laboratory, Central Wharf, Boston, MA 02110
RODHOUSE, Paul, The Laboratory, Marine Biological Association,

Citadel Hill, Plymouth, England PL1 2PB
RODRIGUEZ, Gustov A., PRODEMEX. Apartado Postal 1095,

Los Mochis, Sinaloa, Mexico
ROGERS, Bruce A., 61 Switch Rd., RFD, Hope Valley, RI 02832
ROOSENBURG, Willem H.. Box 16A. Bowen Road, St. Leonard,

MD 20685
ROPER, Dr. Clyde F. E., Dept. of Invertebrate Zoology, Museum of

Natural History, Smithsonian Institution, Washington, DC 20560
ROPES, John W., 21 Pattee Rd., East Falmouth, MA 02536
ROSENBERRY, Robert, 1 1057 Negley Ave., San Diego, CA 92131
ROSENFIELD, Dr. Aaron. National Marine Fisheries Service.

Oxford, MD 21654
ROWELL, Terence W., Fisheries and Oceans, P.O. Box 550, Halifax,

Nova Scotia, Canada B3J 2S7
RUPRIGHT, Gregory L., c/o Sondrini, 7200 Ulmerton Rd., Largo,

FL 33541
RUSSELL, Peggy Rochelle, N 34671 Hwy. 101, Lilhwaup, WA 98555
RYTHER, Dr. John H., Center for Marine Biotechnology, RR1,

Box 196A, Ft. Pierce, FL 33450

SANDIFER, Dr. Paul A., Marine Resources Research Institute,

P.O. Box 12559, Charleston, SC 29412
SAVAGE, Neil, 15 Allen St., Exeter, NH 03833
SAXBY, D. J., 4727 S. Piccadilly, W. Vancouver, British Columbia,

Canada V7W 1J8
SCARPA, John, College of Marine Studies, 700 Pilottown Rd.,

Lewes, DE 19958
SCHILLING, Mary, Harbor Branch Institution Inc., RR1, Box 196A,

Fort Pierce, FL 33450
SCHLICHT, Dr. Frank G., 6711 Rowell Court, Missouri City, TX

77489
SCHNEIDER, R. Randall, Dept. of Natural Resources, Tidewater

Admin., Tawes State Office Bldg. C-2, Annapolis, MD 21401
SCOTT, Timothy, Dept. of Ecology and Evolution, State Univ. of

New York, Stony Brook, NY 11794
SCRO, Robert, New Jersey Dept. of Environmental Protection,

Div. of Water Resources, 25 Arctic Parkway, Trenton, NJ 08625

214

Membership List - National Shellfisheries association

SEGER, James L., 3245 SW Marigold. Portland. OR 97219

SEKI. Tetsuo, Oyster Research Institute, 211 Higashi Mohne

Motoyoshi. Miyagi Prefecture, Japan 988-05
SERCHUK, Dr. Fredric M., National Marine Fisheries Service,

Northeast Fisheries Center. Woods Hole, MA 02543
SHABMAN, Dr. Leonard, Dept. of Agricultural Economics, Virginia

Polytechnic Institute, Blacksburg. VA 24061
SHAW, Harry L., Pacific Aquaculture. P.O. Box 55, Edgecliff,

Sydney, New South Wales 2027 Australia
SHAW, William, Humboldt State Univ., Marine Laboratory, P.O.

Box 624, Trinidad, CA 95570
SHIPMAN, Susan, Dept. of Natural Resources, 1200 Glynn Ave.,

Brunswick, GA 31523
SHIRAISHI, Dr. Kagehide, Dept. of Biology, Iwate Medical Univ.,

Morioka Iwate-Ken. Japan
SHOTWELL, J. A., P.O. Box 417, Bay Center, WA 98527
SHULTZ, Dr. Fred T., P.O. Box 313, Sonoma, CA 95476
SHUMAN, Randy, Applied Marine Research, Inc.. 7525 44th Ave.,

NE, Seattle, WA 98115
SHUMWAY, Dr Sandra, Dept. of Marine Resources, West Boothbay

Harbor, ME 04575
SHUSTER, Dr. Carl N., 3733 N. 25th Street. Arlington, VA 22207
SIDDALL, Dr. Scott E., Marine Sciences Research Center, State

Univ. of New York, Stony Brook, NY 11794
SIEGFRIED, Carol, Univ. of Delaware, 700 Pilottown Rd., Lewes,

DE 19958
SIELING, Fred W., 14 Thompson St., Annapolis. MD 21401
SIELING, F. William, 26 Farragut Rd., Annapolis, MD 21403
SILVIA, Robert, 29 Tri-Town Circle, Mashpee, MA 02649
SIMONS, Donald D., Washington Dept. of Fisheries, 331 State

Highway 12, Montesano, WA 98563
SISSENWINE, Michael P., P.O. Box 12, Woods Hole, MA 02543
SLOAN, Norman A., Pacific Biological Station, Fisheries and

Oceans, Nanaimo, British Columbia, Canada V9R 5K6
SMITH, Bruce W., Public Service Co. of New Hampshire. 1000 Elm

Street, Manchester, NH 03105
SMITH, Dr. John M., Grays Harbor College. Aberdeen, WA 98520
SMITH, Kathleen A.. 396 Appleton St., Arlington, MA 02174
SMITH, Lorene E., Dept. of Biological Sciences, University of New

Orleans, Lakefront, New Orleans, LA 70148
SMITH, Myron C, Coast Oyster Co., P.O. Box 327, Quilcene, WA

98376
SMITH, Theodore I. J., Marine Resources Research Institute, 217 Ft.

Johnson Rd., Charleston, SC 2941 2
SMITH, Walter L., Box 754, Orient, NY 11957
SNYDER, Barry J., Marine Sciences Research Center, State Univ.

of New York, Stony Brook, NY 1 1 794
SONIAT, Dr. Thomas. Dept. of Biological Sciences, Univ. of New

Orleans, Lakefront, New Orleans, LA 70148
SPARKS, Dr. Albert K., Northwest Fisheries Center, 2725 Montlake

Blvd. E, Seattle, WA 98112
STAINKEN, Dennis, 1 Estel Place, Green Brook.NJ 08812
STANLEY, Dr. Jon G., Dept. of Zoology. Univ. of Maine, Orono,

ME 04469
STARR, Richard M., Oregon Dept. of Fish and Wildlife, Bldg. 3,

Marine Science Drive. Newport, OR 97365
STEELE, Eail N., Box 42, Blanchard, WA 98231
STEVENS, Fred S., Marine Resources Research Institute, P.O.

Box 12559. Charleston, SC 29412
STEVENS, Stuart A.. Shellfish Sanitation Program, 1200 Glynn

Ave., Brunswick, GA 31520
STEVENS, Ted, Waddell Mariculture Center, P.O. Box 809, Bluffton,

SC 29910
STEWART, Lance L., Marine Science Institute, Avery Point, Univ.

of Connecticut, Groton, CT 06340

STRASDINE, Susan A.. Institute of Animal Resource Ecology,

Univ. of British Columbia, 2204 Main Mall. Vancouver, British

Columbia, Canada V6T 1W5
STRONG, Craig E., Bluepoints Co., Inc., Foot of Atlantic Ave.,

West Sayville, NY 11796
SUMNER, C. E., 18 Thomas St., North Hobart. Tasmania 7000

Australia
SUNDERLIN, Judith B.. 58E Cotton Valley, Star Rt. 00864,

Christiansted, St. Croix, Virgin Islands 00820
SUPAN, John, Fisheries Agent, Cooperative Extension Service,

P.O. Box 2440, Covington, LA 70434
SUPRENANT, Albert H., Cape Cod Oyster, 262 Bridge St., Oster-

ville, MA 02655
SWAN, William H., P.O. Box 758, Hampton Bays, NY 11946
SWEENY, Brian, P.O. Box 914, Gloucester Point, VA 23062
SWIFT, Dr. Mary L., 15656 Millbrook Lane, Laurel, MD 20707
SZIKLAS, Robert, Wauwinet, Nantucket, MA 02554

TABARINI, C. L. 7836 Midday Lane, Alexandria, VA 22306
TAUB, Dr. Frieda B., College of Fisheries. Univ. of Washington,

Seattle, WA 981 95
TAYLOR, David M., P.O. Box 5667, St. John's Newfoundland.

Canada A 1C 5X1
TAYLOR, Frank S., Marine Resources Research Institute, P.O.

Box 12559, Charleston, SC 29412
TAYLOR, Rodman E., Shellfish Unit, School of Fisheries WH-10,

Univ. of Washington, Seattle, WA 98195
TEMPLETON, Dr. James E., c/o W & P Nautical, Inc. 222 Severn

Ave., Annapolis. MD 21403
TETTELBACH, Lisa Petti. National Marine Fisheries Service,

212 Rogers Ave. .Milford, CT 06460
TETTELBACH, Stephen, Marine Research Laboratory, Univ. of

Connecticut, Noank, CT 06340
THOMAS, Dr. M. L. H., Dept. of Biology, Univ. of New Brunswick,

P.O. Box 5050, St. John, New Brunswick, Canada E2L 415
THOMPSON, Douglas S.. P.O. Box 196, Nanoos Bay, British

Columbia, Canada V0R 2R0
THOMPSON, Richard, 2902 Dillionhill Drive, Austin, TX 78745
THURBERG, Dr. Frederick P., National Marine Fisheries Service,

212 Rogers Ave., Milford, CT 06460
TOLL, Dr. Ronald B., Dept. of Biology. Univ. of the South,

Sewanee, TN 37375
TOLLEY, Everett A.. Progressive Services Inc., P.O. Box 10076,

Baltimore, MD 21204
TOWNSHEND, E. Roger, Blooming Point Rd., Rural Route 1,

Mount Stewart, Prince Edward Island, Canada C0A 1T0
TREVELYAN, George, Univ. of California, Bodega Marine Lab.,

P.O. Box 247, Bodega Bay, CA 94923
*TRUITT, Dr. Reginald V, Great Neck Farm, Stevensville, MD 21666
TURNER, Dr. Ruth D., Museum of Comparative Zoology, Harvard

Univ., Cambridge, MA 02138
TWEED, Stewart, Cape May County Extension Office. Dennisville

Road, Rte. 657, Cape May Court House, NJ 08204

UKELES, Dr. Ravenna, National Marine Fisheries Service, 212

Rogers Ave., Milford, CT 06460
URBAN, Edward R., College of Marine Studies, Univ. of Delaware,

Lewes, DE 19958

VACAS, Lie. Herman C, Estacion Pesquera Experimental, Avda.

Costanera, 8520 San Antonia Oeste, Reo Negro, Argentina
VAN ENGEL, Willard A., Virginia Institute of Marine Science,

College of William and Mary, Gloucester Point, VA 23062
VAN HEUKELEM, Dr. William F., Horn Point Environmental Lab.,

Univ. of Maryland, P.O. Box 775, Cambridge, MD 21613

Membership List - National Shellfisheries association

215

VAN VOLKENBURGH, Pieter, 464 Greene Ave. , SayviUe, NY 1 1 782
VAUGHAN, David E., 159 Flamingo Rd., Tuckerton, NJ 08087
VELEZ, Anibal, Instituto Oceanographico, Apartado Postal 308,

Cumana 6101 Venezuela
VERGARA, Victor M., 7622 Democracy Blvd., Bethesda, MD 21817
VOLK, John H., Dept. of Agriculture, Aquaculture Div., P.O.

Box 97, Milford, CT 06460
VOUGLITOIS, James J., G.P.U. Nuclear Environmental Control,

P.O. Box 388, U.S. Route 9, Forked River, NJ 08731

WADA, Katsuhiko, National Research Institute of Aquaculture,

Nansei, Mie 516-01 Japan
WAGNER, Eric, 1632 Mayfair Ct., Point Pleasant, NJ 08742
WALKER, Randal L.. Skidaway Institute of Oceanography, P.O.

Box 13687, Savannah, GA 31416
WALLACE, Dana E., 3081 Mere Pt. Road, Brunswick, ME 0401 1
WALLER, Dr. Thomas R., Curator, Dept. of Paleobiology, Smith-
sonian Institution. Washington, DC 20560
WALSH, Dennis T., Aquaculture Research Corp., P.O. Box 597,

Dennis, MA 02368
WALSH, M. G., Dept. of Bioresource Engineering, Univ. of British

Columbia, Vancouver, British Columbia, Canada
WARD, Jonathan Evan, CR 3, Box 1059, Lewes, DE 19958
WATSON, R. H., P.O. Box 876, Bicheno, Tasmania 7215 Australia
WAUGH, Godfrey R., Wallace Groves Aquaculture Foundation,

P.O. Box 140939, Coral Gables, FL 33114
WEIL, Ernesto, Fundacion Cientifica Los Roques, Apartado 1

Carmelitas, Caracas 1010-A Venezuela
WEINER, Ronald, Microbiology Dept., Univ. of Maryland, College

Park, MD 20742
WEINHEIMER, Debra Ann, National Marine Fisheries Service,

Ft. Johnson Road, Charleston, SC 29407
WEISS, Prof. Charles M., Dept. of Environmental Sciences and

Engineering, Univ. of North Carolina, 104 Rosenau Hall, Chapel

Hill, NC 27514
WENNER, Dr. Elizabeth Lewis, South Carolina Marine Resources

Research Institute, P.O. Box 12559, Charleston, SC 29412
WESTLEY, Ronald E., 6606 Sierra Dr., SE, Lacey, WA 98503
WHEATON. Dr. Fred, Univ. of Maryland, Dept. of Agricultural

Engineering, College Park, MD 20742
WHITAKER, J. David, South Carolina Wildlife and Marine Resources

Div., P.O. Box 12559, Charleston, SC 29412

WHITCOMB, James P., Star Route Box 35, Gloucester Point, VA

23062
WHITE, Marie, Dept. of Oceanography, Texas A&M Univ., College

Station, TX 77843
WIDMAN, James, National Marine Fisheries Service, Northeast

Fisheries Center, Milford, CT 06460
WIKFORS, Gary H., National Marine Fisheries Service Laboratory,

212 Rogers Ave., Milford, CT 06460
WILEY, Cloyde W., Rte 2, Box 65, Quinton, VA 23141
WILLIS, Scott, Florida Dept. of Natural Resources, Marine Research

Lab., 100 8th Avenue SE, St. Petersburg, FL 33701
WILSON, Kerry A., New Brunswick Dept. of Fisheries. P.O. Box

6000, Fredericton, New Brunswick, Canada E3B 5H1
WINSTANLEY, Ross H., Comm. Fisheries Branch, Fisheries &

Wildlife Service, 250 Victoria Parade, P.O. Box 41, East

Melbourne, Australia 3002
WOELKE, Dr. Charles, Washington Dept. of Fisheries, General

Administration Bldg., Olympia, WA 98501
WOLF, Peter H., 62 Mackenzie St., Bondi Junction. New South

Wales 2022 Australia
WOON, Gail L., Center for Marine Biotechnology, Harbor Branch

Institution, Rt. 1, Box 196A, Ft. Pierce, FL 33450

YOUNG, Adam, Seafarming Project, SEAFDEC, P.O. Box 256,

Iloilo City, Philippines 5901
YOUNG, Brenda L., Dept. of Biology, Univ. of South Carolina,

Columbia, SC 29208
YOUNG, James S., Battelle Marine Research Laboratory. 439 W.

Sequim Bay Rd., Sequim, WA 98382
YOUNG, Jeffrey, Pacific Seafood Industries, P.O. Box 2544,

Santa Barbara, CA 93120

ZAHTILA, Joseph J., 122 Bayville Ave.. Bayville, NY 11709
ZIMMERMAN, John M., 122 Hoyt St., Apt. IE, Stamford, CT

06905
ZOTO, Dr. George A., 10 Widgeon Lane, West Barnstable, MA

02668

PACIFIC COAST SECTION
NATIONAL SHELLFISHERIES ASSOCIATION

(As of 1 January 1984)

ALLEN, Stan, School of Fisheries WH-10, Univ. of Washington,

Seattle, WA 98195
AMPAK, 9451 A Van Home Way, Richmond, British Columbia.

Canada V6X 1W2
ANDERSON, Greg, 1572 River Rd., Brunswick, ME 0401 1
ARMSTRONG, Dr. David A., School of Fisheries WH-10, Univ. of

Washington, Seattle, WA 98195
ARMSTRONG, John W., 6045 NE 51st, Seattle, WA 981 15

BALDASSCM, Brian, 1619 N. Warner. No. 2. Tacoma, WA 98407
BATCHELDER, Jack, Coast Oyster Co., P.O. Box 327, Quilcene,

WA 98376
BAYNES Sound Oyster Co., Ltd., P.O. Box 127, Union Bay.

British Columbia, Canada V0R 3B0
BEATTIE, J. Hal, National Marine Fisheries Service Aquaculture

Station. Univ. of Washington Facility, P.O. Box 38, Manchester,

WA 98353
BEAUDRY, Jerry, School of Fisheries WH-10, Univ. of Washington,

Seattle, WA 98195

BETTINGER, Tom, Washington State Shellfish Lab.. 1000 Pt.

Whitney Rd., Brinnon, WA 98320
BOHN, Richard, Wiegardt & Sons, Inc., P.O. Box 189, Ocean Park,

WA 98640
BONACKER, Gregg, 4033 Corliss Ave. N., Seattle, WA 98103
BOULE, Marc, Shapiro and Assoc, 1812 Smith Tower, Seattle,

WA 98104
BOURNE, Dr. Neil, Pacific Biological Station, Nanaimo, British

Columbia, Canada V9R 5K6
BREEN, Paul, Fisheries Canada, Pacific Biological Station. Nanaimo,

British Columbia, Canada V9R 5K6
BREESE, Prof. Willy, Oregon State Univ., Marine Science Center,

Newport, OR 97365
BRONSON, Jeff, Shellfish Laboratory, 1000 Pt. Whitney Rd..

Brinnon, WA 98320
BROWN, Jim, Dept. of Biological Sciences, Simon-Fraser Univ.,

Burnaby, British Columbia. Canada V5A 1S6
BUDNICK, Nicholas D., Consolidated Net & Twine Co., 1549 NW

49th St., Seattle, WA 98107

216

Membership List - National Shellfisheries Association

BURBANK, Christine, Coast Oyster Co., P.O. Box 327, Quilcene.

WA 98376
BURGE, Richard, Washington State Dept. of Fisheries, 1000 Pt.

Whitney Rd., Brinnon, WA 98320

GLUDE, John, 2703 W. McGraw, Seattle, WA 98119
GOODWIN, Lynn, Rt. 2, Box 711, Quilcene, WA 98376
GR1SCHKOWSKY, Dr. Roger, Alaska Dept. of Fisheries, 333 Rasp-
berry Rd., Anchorage, AK 99502

CALOMENI, Dave, North Seattle Community College, Biology

Dept., 9600 CoUege Way N., Seattle, WA 98103
CAMPBELL-ATHERTON, Moira, 259 Northridge Dr.. Shawano,

WI 54166
CAMPBELL, Virginia, 1177 Forge Walk, Vancouver, British

Columbia, Canada V6P 3R1
CANADIAN Benthic Ltd., P.O. Box 97, Bamfield, British Columbia,

Canada V0R 1B0
CARRASCO, Ken, School of Fisheries WH-10. Univ. of Washington,

Seattle, WA 98195
CARDWELL, Rick D.. Envirosphere Co., 400-112th Ave. NE.

BeUevue, WA 98004
CASIMIR, Al, 2616 Kwina Rd., Bellingham, WA 98225
CHEW, Dr. Kenneth K., School of Fisheries WH-10, Univ. of Wash-
ington, Seattle, WA 98195
CLELAND, Bill. 1604 N. Bethel, Olympia, WA 98506
CONTE, Dr. Fred, Aquaculture Extension, University of California,

Davis, CA 95616
COOPER, Ken, Shellfish Laboratory, 1000 Pt. Whitney Rd., Brinnon,

WA 98320
COX, Robert, Marine Resources Branch, Parliament Bldgs., Victoria,

British Columbia, Canada V8V 1X5
CREEKMAN, Laura, Washington State Dept. of Fisheries, P.O. Box

190, Ocean Park, WA 98640
CUMMINS, Joseph M., Environmental Protection Agency, Box 549,

Manchester, WA 98353
CUDD, Sue, 2809 165th Place, NE, Bellevue, WA 98008

DAVIS, Joth, School of Fisheries WH-10, Univ. of Washington,

Seattle, WA 98195
De MARTINI, Dr. John, 1111 Birch Ave., McKinleyville, CA 95521
DEMORY, Darrell, Oregon Dept. of Fish & Wildlife, Marine Science

Drive, Newport, OR 97365
DONALDSON, James, Coast Oyster Co., P.O. Box 327, Quilcene,

WA 98376
DRISCOLL, John M., Northwestern Glass Co., 20065 SW Santa

Rosa Ct., Beaverton. OR 97007
DUNGAN, Christopher, 10021 NE 122nd, No. D, Kirkland, WA

98034

ECHOLS, Louie S., Director, Washington Sea Grant Program,
3716 Brooklyn Ave. NE, Seattle, WA 98195

ELSTON, Dr Ralph, Senior Research Scientist. BatteUe North-
west Div., 439 W. Sequim Bay Rd., Sequim, WA 98382

EMMETT, Brian, P.O. Box 6418 Station 'C, Victoria, British
Columbia, Canada J8P 5M3

ERVEST, Mrs. Ray, Salty Dog Seafood, 5823 Steamboat Island Rd.,
Olympia, WA 98502

FALMAGNE, Catherine, 16718 76th Ave. NE, Bothell, WA 98011
FAUDSKAR, John, Oregon State Univ. Extension Service, 2204

4th St., Tillamook, OR 97141
FOLLETT, Jill, 10300 Schneiter Dr., Anchorage, AK 99516
FOSTER, Carolyn, Biological Structure SM-20, Univ. of Washington,

Seattle, WA 98195
FULLER, Julie, 919 NE 71st, Seattle, WA 98117

GANGMARK, Carolyn, Environmental Protection Agency Labora-
tory, P.O. Box 549. Manchester, WA 98353
GIORGI, Al, 812 NE 83rd, Seattle, WA 98115

HAARS, Ellen, Dept. of Social & Health Services, MSLD 11,

Olympia, WA 98502
HALL, Sherwood, WHOl/Clark 410, Woods Hole, MA 02543
HANSON, Leigh, 9705 N. Edison, Portland, OR 97203
HAZELTINE, Arthur, Marine Culture Lab., Coast Route, Granite

Canyon, Monterey, CA 98940
HAYS, Max G, Dept. of Social & Health Services, Div. LD-11,

Olympia, WA 98504
HERITAGE, Dwight. Fisheries & Environment. Pacific Biological

Station, Box 100, Nanaimo, British Columbia, Canada V9R 5K6
HERSHBERGER, Dr. William K., School of Fisheries WH-10,

Univ. of Washington, Seattle, WA 98195
HOFFMAN, Ethelyn G, E. 1261 Mason Lake Dr. E., Grapeview,

WA 98546
HOGG Island Oyster, 127V2 Darwin St., Santa Cruz, CA 95060
HOWLAND, Paul 283 Old Blyn Hwy„ Sequim. WA 98382
HUMPHREYS, Jim, Washington Sea Grant, 19 Harbor Mall. Belling-
ham, WA 98225
HURLBURT, Eric, 1412 NW 61st, Seattle, WA 98107

IM, Kwang H., c/o Earl R. Combs, Inc., 9725 SE 36 th, Mercer Island,

WA 980404
INCZE, Dr. Lewis, School of Fisheries WH-10. Univ. of Washington,

Seattle, W A 98195
IWAMOTO, Robert N., School of Fisheries WH-10, Univ. of

Washington, Seattle, WA 98195

JAMBOR, Nick, Box 465 Star Rt., South Bend, WA 98586
JEFFERDS, Peter, Penn Cove Mussels, P.O. Box 148, Coupeville,

WA 98239
JOHNSON, Kurt, 1515 NE 105th, Seattle, WA 98125
JONES, Bruce and Gordon, Innovative Aquaculture Ltd., Skerry

Bay, Lasqueti Island, British Columbia, Canada V0R 2S0

KELLY, Randolph O, 1234 E. Shaw, Fresno, CA 93710
KLINE, Thomas, 1725 NE 90th, Seattle, WA 981 15
KU1PER, Ted, 912 K St., Eureka, CA 95501
KYTE, Michael, 527 212th SW, Bothell, WA 9801 1

LAGOON Seafoods Ltd., 1317 Walnut St., Vancouver, British

Columbia, Canada V6S 3R2
LANGMO, Don, Industrial Engineer, Dept. of Agricultural and

Resource Economics, Oregon State Univ., Corvallis, OR 97331
LILJA, Jack, Dept. of Social & Health Services Div., LD-11,

Olympia. WA 98504
L1MBERIS Seafoods Ltd., Box 568, Ladysmith, British Columbia,

Canada V0R 2E0
LINDSEY, Cedric E., 744 Pt. Whitney Rd., Brinnon, WA 98320
LIPOVSKY, Vance & Sandy, P.O. Box 635, Ocean Park. WA 98640
LOOSANOFF, Dr. Victor, 17 Los Cerros Dr., Greenbrae. CA

94902
LOTOSKI, Doug, 181 Citrus Ave., Imperial Beach, CA 92032

MADENWALD, Darlene, Western Washington Univ., Shannon Point

Marine Laboratory, Anacortes, WA 98221
MAGOON, Doug, Dept. of Natural Resources, EK-12, Olympia,

WA 98504
MANAHAN, Dr. Donald, Dept. of Biology, Univ. of Southern Calif,

Los Angeles, CA 90089

Membership List - National Shellfisheries association

217

MARTIN, Roy E., National Fisheries Institute, 2000 M St. NW,

Washington, DC 20036
MATTHEWS, Robert, P.O. Box 494, Ocean Park, WA 98640
McGRAW, Dr. Kay, 131 N. 4th, Seattle, WA 98103
MEURER, David A., Buckhorn Inc., 1690 Naomi Ct., Redwood

City, CA 9406 1
MILLER, Mark B.. 2526 State St., Everett, WA 98201
MIX, Michael C, Dept. of General Sciences. Oregon State Univ.,

Corvallis, OR 97331
MUELLER, Bernie, Marrostone Oysters, 3121 Flagler Rd., Nord-

land, WA 98358
MUMAW, Laura, Seattle Marine Aquarium, Pier 59, Seattle, WA

98191
MUSGROVE, Nancy, 6538 Earl Ave. NW, Seattle, WA 98117

NAKATANI, Dr. Roy E., School of Fisheries WH-10, Univ. of
Washington, Seattle, WA 98195

NEVE, Dr. Richard, Institute of Marine Sciences, Univ. of Alaska.
Fairbanks, AK 99701

NISHITANI, Louisa, School of Fisheries WH-10. Univ. of Wash-
ington, Seattle, WA 98195

NORTHRUP, Tom, Washington State Dept. of Fisheries, Coastal
Research Shellfish Labs, 331 State Highway 12, Montesano.
WA 98563

NOSHO, Terry, Washington Sea Grant Office, Univ. of Washington,
3716 Brooklyn NE. Seattle, WA 98195

OLIVER, Susan, 5390 Schmitt Rd., Port Angeles, WA 98362
OLSEN, Scharleen, Washington Dept. of Fisheries, 1000 Pt. Whitney

Rd., Binnon, WA 98320
OSIS, Laimons. Oregon Dept. of Fish & Wildlife, Bldg. 3, Marine

Science Dr., Newport, OR 97365
OSTASZ, Michael J., State of Alaska, Dept. of Environmental

Conservation, P.O. Box 10-4240, Anchorage, AK 99510

PCOGA, 1437 Elliott Ave. W.. Seattle, WA 98119

PAMENES, Luis Garcia, Instituto de Invest. Oceano., Univ. Auton.

de Baja California, Apartado Postal 453, Ensenada, Baja

California, Mexico
PERDUE, James, 1709 Upper Millstone Ln.. Salisbury, MD 21801
PETERS, John B., Univ. of Washington, Washington Sea Grant

Program HF-10, Seattle, WA 98195
POPHAM, Dr. David. Seakem Oceanography Lot, 2045 Mills Rd.,

Sidney, British Columbia, Canada V8L 3S1
PRENTICE, Earl, Fisheries Res. Biol., National Marine Fisheries

Service, P.O. Box 38, Manchester, WA 98353

QUAYLE, Dr. Daniel B., Pacific Biolgoical Station, Nanaimo,
British Columbia, Canada V9R 5K6

RAVEN, Gary, P.O. Box 783, Coupeville, WA 98239
RENSEL, Jack. 8249 Corliss Ave. N„ No. 2, Seattle, WA 98103
ROBINSON, Anja, Oregon State Univ., Marine Science Center,

Newport, OR 97365
ROSENBERRY, Robert, Aquaculture Digest, 9434 Kierny Mesa

Rd, San Diego, CA 92126

ROTEN, Dr. Robert, Aquatic Research Inst. 2242 Davis Ct.,

Hayward, CA 94545
RUSH, Ralph, Marrowstone Oyster Farm, 3081 Flagler Rd.,

Nordland, WA 98358

SCHOLZ, Al, Washington Dept. of Fisheries, P.O. Box 224,

Quilcene, WA 98376
SEYMOUR, Steve, 2616 Kwina Rd., Bellingham, WA 98226
SHRINER, Jan, P.O. Box 93, Brinnon, WA 98320
SIMON, Doug, Washington Dept. of Fisheries, Coastal Shellfish

Lab., Montesano. WA 98563
SK1DMORE, Doug, P.O. Box 783, Coupeville, WA 98239
SMITH, Myron C, Coast Oyster Co., P.O. Box 327, Quilcene,

WA 98376
SPARKS, Dr. Albert, National Marine Fisheries Service, 2725

Montlake Blvd. E.. Seattle, WA 98112
STACY, Robert, 14541 SE 167th, Renton, WA 98055
STEELE, Earl, 730 Old 99 N., Burlington, WA 98233
STERN, Roger, 10778 NE Seaborn Rd., Bainbridge Island, WA

98110
SYNDEL Laboratories, Ltd.. 8879 Selkirk St., Vancouver, British

Columbia, Canada V6P 4S6

TAUB, Dr. Freida, School of Fisheries WH-10, Univ. of Washington,

Seattle, WA 98195
TAYLOR, Rodman, School of Fisheries WH-10, Univ. of Wash-
ington, Seattle, W A 98195
THOMPSON, Doug, Pacific Biological Station, Nanaimo, British

Columbia, Canada V9R 5K6
TOKAR, Erick M., ITT Rayonier Research Center, 409 E. Harvard,

Shelton, WA 98584
TOLLEY, Everett A., President, Progressive Services, Inc., P.O.

Box 10076, Baltimore, MD 21204
TUFTS, Dennis, Washington State Dept. of Fisheries, P.O. Box 190,

Ocean Park, WA 98640
TYNAN, Tim, Squaxin Indian Tribe, W81 Highway 108, Shelton,

WA 98584

VAN CITTERS, Bob, 14630 Norma Beach Rd., Edmonds, WA
98020

WACHSMITH, Lou, Oregon Oyster Co., 208 SW Ankeny St.,

Portland, OR 97204
WARING, Arnold, 1437 Elliott Ave., Seattle, WA 98119
WATERSTRAT, Paul, Drawer V, Mississippi State Univ., Mississippi

State, MS 39762
WATSON, Bob, c/o IAP, Ltd., Skerry Bay. Lasquiti Island, British

Columbia, Canada V0R 2J0
WESTLEY, R. E., 6606 Sierra Dr. SE, Lacey, WA 98504
WOELKE, Dr. Charles, Washington State Dept. of Fisheries, 2378

Crestline Blvd., Olympia, WA 98502

YAMASHITA, Jerry. Western Oyster Co., 920 E. Allison, Seattle,
WA 98102

ZAHRADINK, John W.,7771 Ash St., Richmond, British Columbia,
Canada V6Y 2S2

218

Membership List - National Shellfisheries Association

SUBSCRIBING INSTITUTIONS

(As of 1 January 1984)

AUSTRALIA

New South Wales

Div. of Fisheries & Oceanography, CSIRO Library, P.O. Box 21,

Cronulla, New South Wales, Australia 2230
New South Wales State Fisheries, 211 Kent St. (Fisheries House),

Sydney, New South Wales, Australia 2000

Queensland

The Librarian, Queensland Fisheries Service, P.O. Box 344, Fortitude

Valley, Queensland, Australia 4006
The Library/Serials Post Office, lames Cook Univ., Queensland,

Australia 48 1 1

South Australia

R. Hill & Son, Ltd.. Subscription Agents, 29 King William St.,
Adelaide. South Australia, Australia 5000

West Australia

Librarian (2096/71-72), Dept. Fisheries & Wildlife, 108 Adelaide
Terrace, Perth, West Australia, Australia 6000

BELGIUM

W. H. Smith & Son, 71 Blvd. Adolphe Max, 1000 Brussels, Belgium

CANADA

British Columbia

Fisheries & Oceans Library, Pacific Biological Station, P.O. Box 100,

Nanaimo, British Columbia, Canada V9R 5K6
IDRC Library, Univ. of British Columbia, 5990 Iona Dr., Vancouver.

British Columbia, Canada V6T 1 L4
Redonda Sea Farms. Ltd., Refuge Cove, British Columbia, Canada

V0P 1P0
Woodward Biomedical Lab., Serials Div., Univ. of British Columbia,

2198 Health Sciences Mall, Vancouver, British Columbia, Canada

V6T 1W5

New Brunswick

Dept. of Fisheries & Oceans, Biological Station Library, St. Andrews.

New Brunswick, Canada E0G 2X0
Fisheries & Oceans Library, Atlantic Fisheries Gulf Region, P.O.

Box 5030, Moncton, New Brunswick, Canada E1C 9B6

Newfoundland

Librarian, College of Fisheries, P.O. Box 4920, St. John's,

Newfoundland, Canada A1C 5R3
Periodical Div. (P-35454), Main Library, Memorial Univ. of

Newfoundland, Canada A1B 3Y1
Regional Library, Fisheries & Oceans Canada, NWAFS, P.O. Box

5667, St. John's, Newfoundland, Canada A1C 5X1

Nova Scotia

Canada Fisheries & Oceans, Scotia Fundy Library, P.O. Box 550,

Halifax, Nova Scotia, Canada B3J 2S7
Killam Library Serials Dept., Dathousie Univ., Halifax, Nova Scotia,

Canada B3H 4H8

Ontario

Fisheries & Oceans Library, Ottawa, Ontario, Canada K1A 0E6
LSI 4039, Canada Inst, for S.T.I. , Library Serials Acquisitions,
National Research Council, Ottawa, Ontario, Canada K1A 0S2

Quebec

Div. Des Acquisitions Periodiques, 510444 Bibl. de L'Universite
Laval, Quebec, Quebec, Canada G1K 7P4

CHILE

Library, Inst, de fomento Pesquero, Av. Pedro de Valdivia 2633,

Santiago, Chile
Univ. del Notre Biblioteca, Avda Angamos 0610, Casilla No. 1280,

Antofagasata, Chile

FEDERAL REPUBLIC OF GERMANY
(WEST GERMANY)

Biologische Anstalt Helgoland, Bibliothek, Notkestrasse 31, D-2000,

Hamburg 52, West Germany
Institut fur Meeresforschung Bibliothek, Am Handelshafen 12,

D-2850 Bremerhaven 1, West Germany
Staats und Universitatsbibliothek, ABT-DF, Von-Melle-Park 3. 2000

Hamburg 13, West Germany
Th. Christiansen Bookseller, Bahrenfelder Str. 79, Postif. 5 03 06,

2000 Hamburg 50 (Altona), West Germany

HOLLAND

Rijksinstituut voor Visserijonderzoek, Postbus 68, 1970 AB
Ymuiden, Holland 1

HONG KONG

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JAPAN

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Rovinj, Yugoslavia

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JOURNAL OF SHELLFISH RESEARCH

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JOURNAL OF SHELLFISH RESEARCH
Vol. 3, No. 2 December 1983

CONTENTS

Edwin W. Cake, Jr.

Symbiotic Associations Involving the Southern Oyster Drill Thais haemastoma floridana

(Conrad) and Macrocrustaceans in Mississippi Waters 117

Robert W. Elner and Rene E. Lavoie

Predation on American Oysters (Crassostrea virginica [Gmelin] ) by American Lobsters
(Homarus americanus Milne-Edwards), Rock Crabs (Cancer irroratus Say), and Mud
Crabs [Neopanope sayi [Smith] ) 129

M. F. Li, R. E. Drinnan, Michael Drebot, Jr. and Gary Newkirk

Studies of Shell Disease of the European Flat Oyster Ostrea edulis Linne in Nova Scotia 135

Dexter S. Haven and James P. Whitcomb

The Origin and Extent of Oyster Reefs in the James River, Virginia 141

Norman E. Buroker

Genetic Differentiation and Population Structure of the American Oyster Crassostrea

virginica (Gmelin) in Chesapeake Bay 153

Randal L. Walker

Feasibility of Mariculture of the Hard Clam Mercenaria mercenaria Linne in Coastal Georgia 169

Don P. Man the, Ronald F. Malone and Harriet M. Perry

Water Quality Fluctuations in Response to Variable Loading in a Commercial, Closed

Shedding Facility for Blue Crabs '. 1 . .'". : . u '.. ': .' 175

George R. Abbe

Blue Crab (Callinectes sapidus Rathbun) Populations in Mid-Chesapeake Bay in the

Vicinity of the Calvert Cliffs Nuclear Power Plant, 1968-1981 183

..... .• ■ •• ■'• • ■- '"''

William E. Donaldson "..._..• '

Movements of Tagged Males of Tanner Crab Chionoecetes bairdi Rathbun off Kodiak Island, Alaska 195

RESEARCH NOTE

M. C Gibbons, J. G. Goodsell, M. Castagna and R. A. Lutz

Chemical Induction of Spawning by Serotonin in the Ocean Quahog A re tica islandica (Linne) 203

Membership Listing of the National Shellfisheries Association 207

COVER PHOTOGRAPH (1.5 X): Florida rock-shell Thais haemastoma floridana (Conrad), also known as
the "southern oyster drill," on shell of the eastern oyster Crassostrea virginica (Gmelin). Note the drill hole on
the small attached oyster. Specimens were collected from Biloxi Bay, Mississippi. [Photograph taken by
Dr. Robin Overstreet and printed by Joan Durfee, Gulf Coast Research Laboratory, Ocean Springs, MS.]

MBI. WHOI LIBRARY

UH lAAb X