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Full text of "The Biological bulletin"

Volume 173 



Number 1 



THE 



BIOLOGICAL BULLETIN 



PUBLISHED BY 

THE MARINE BIOLOGICAL LABORATORY 
Editorial Board 



RUSSELL F. DOOLITTLE, University of California at 

San Diego 

WILLIAM R. ECKBERG, Howard University 
ROBERT D. GOLDMAN, Northwestern University 

C. K. GOVIND, Scarborough Campus, University 

ofToronto 

JUDITH P. GRASSLE, Marine Biological Laboratory 

MICHAEL J. GREENBERG, C. V. Whitney Marine 
Laboratory, University of Florida 

MAUREEN R. HANSON, Cornell University 
JOHN E. HOBBIE, Marine Biological Laboratory 
LIONEL JAFFE, Marine Biological Laboratory 



WILLIAM R. JEFFERY, University of Texas at Austin 

GEORGE M. LANGFORD, University of North 

Carolina at Chapel Hill 

GEORGE D. PAPPAS, University of Illinois at Chicago 
SIDNEY K. PIERCE, University of Maryland 

HERBERT SCHUEL, State University of New York at 

Buffalo 

VIRGINIA L. SCOFIELD, University of California at 
Los Angeles School of Medicine 

LAWRENCE B. SLOBODKIN, State University of New 

York at Stony Brook 

JOHN D. STRANDBERG, Johns Hopkins University 
DONALD P. WOLF, Oregon Regional Primate Center 



HOLGER W. JANNASCH, Woods Hole Oceanographic SEYMOUR ZIGMAN, University of Rochester 

Institution 



Editor: CHARLES B. METZ, University of Miami 



AUGUST, 1987 



Printed and Issued by 
LANCASTER PRESS, Inc. 

PRINCE &. LEMON STS. 
LANCASTER, PA 



Marine Biological Laboratory 
^LIBRARY 

: - : SEP 141987 ! 



Woods Hole, Mass. 



THE BIOLOGICAL BULLETIN 

THE BIOLOGICAL BULLETIN is published six times a year by the Marine Biological Laboratory, MBL 
Street, Woods Hole, Massachusetts 02543. 

Subscriptions and similar matter should be addressed to THE BIOLOGICAL BULLETIN, Marine Bio- 
logical Laboratory, Woods Hole, Massachusetts. Single numbers, $20.00. Subscription per volume (three 
issues), $50.00 ($100.00 per year for six issues). 

Communications relative to manuscripts should be sent to Dr. Charles B. Metz, Editor, or Pamela 
Clapp, Assistant Editor, at the Marine Biological Laboratory, Woods Hole, Massachusetts 02543. 



POSTMASTER: Send address changes to THE BIOLOGICAL BULLETIN, Marine Biological Laboratory, 

Woods Hole, MA 02543. 

Copyright 1987, by the Marine Biological Laboratory 
Second-class postage paid at Woods Hole, MA, and additional mailing offices. 

ISSN 0006-3 185 



INSTRUCTIONS TO AUTHORS 

The Biological Bulletin accepts outstanding original research reports of general interest to biologists 
throughout the world. Papers are usually of intermediate length (10-40 manuscript pages). Very short 
papers (less than 10 manuscript pages including tables, figures, and bibliography) will be published in a 
separate section entitled "Short Reports." A limited number of solicited review papers may be accepted 
after formal review. A paper will usually appear within four months after its acceptance. 

The Editorial Board requests that manuscripts conform to the requirements set below; those manu- 
scripts which do not conform will be returned to authors for correction before review. 

1. Manuscripts. Manuscripts, including figures, should be submitted in triplicate. (Xerox copies of 
photographs are not acceptable for review purposes.) The original manuscript must be typed in double 
spacing (including figure legends, footnotes, bibliography, etc.) on one side of 16- or 20-lb. bond paper, 8'/2 
by 1 1 inches. Manuscripts should be proofread carefully and errors corrected legibly in black ink. Pages 
should be numbered consecutively. Margins on all sides should be at least 1 inch (2.5 cm). Manuscripts 
should conform to the Council of Biology Editors Style Manual, 4th Edition (Council of Biology Editors, 
1978) and to American spelling. Unusual abbreviations should be kept to a minimum and should be spelled 
out on first reference as well as defined in a footnote on the title page. Manuscripts should be divided into 
the following components: Title page. Abstract (of no more than 200 words). Introduction, Materials and 
Methods, Results, Discussion, Acknowledgments, Literature Cited, Tables, and Figure Legends. In addi- 
tion, authors should supply a list of words and phrases under which the article should be indexed. 

2. Figures. Figures should be no larger than 8'A by 1 1 inches. The dimensions of the printed page, 5 
by 7% inches, should be kept in mind in preparing figures for publication. We recommend that figures be 
about l'/2 times the linear dimensions of the final printing desired, and that the ratio of the largest to the 
smallest letter or number and of the thickest to the thinnest line not exceed 1:1.5. Explanatory matter 
generally should be included in legends, although axes should always be identified on the illustration itself. 
Figures should be prepared for reproduction as either line cuts or halftones. Figures to be reproduced as 
line cuts should be unmounted glossy photographic reproductions or drawn in black ink on white paper, 
good-quality tracing cloth or plastic, or blue-lined coordinate paper. Those to be reproduced as halftones 
should be mounted on board, with both designating numbers or letters and scale bars affixed directly to 
the figures. All figures should be numbered in consecutive order, with no distinction between text and plate 
figures. The author's name and an arrow indicating orientation should appear on the reverse side of all 
figures. 

3. Tables, footnotes, figure legends, etc. Authors should follow the style in a recent issue of The 
Biological Bulletin in preparing table headings, figure legends, and the like. Because of the high cost of 
setting tabular material in type, authors are asked to limit such material as much as possible. Tables, with 
their headings and footnotes, should be typed on separate sheets, numbered with consecutive Roman 
numerals, and placed after the Literature Cited. Figure legends should contain enough information to 
make the figure intelligible separate from the text. Legends should be typed double spaced, with consecutive 
Arabic numbers, on a separate sheet at the end of the paper. Footnotes should be limited to authors' current 
addresses, acknowledgments or contribution numbers, and explanation of unusual abbreviations. All such 
footnotes should appear on the title page. Footnotes are not normally permitted in the body of the text. 



4. A condensed title or running head of no more than 35 letters and spaces should appear at the top of 
the title page. 

5. Literature cited. In the text, literature should be cited by the Harvard system, with papers by more 
than two authors cited as Jones et al., 1980. Personal communications and material in preparation or in 
press should be cited in the text only, with author's initials and institutions, unless the material has been 
formally accepted and a volume number can be supplied. The list of references following the text should 
be headed LITBRATURE CITED, and must be typed double spaced on separate pages, conforming in 
punctuation and arrangement to the style of recent issues of The Biological Bulletin. Citations should 
include complete titles and inclusive pagination. Journal abbreviations should normally follow those of 
the U. S. A. Standards Institute (USASI), as adopted by BIOLOGICAL ABSTRACTS and CHEMICAL AB- 
STRACTS, with the minor differences set out below. The most generally useful list of biological journal titles 
is that published each year by BIOLOGICAL ABSTRACTS (BIOSIS List of Serials; the most recent issue). For- 
eign authors, and others who are accustomed to using THE WORLD LIST OF SCIENTIFIC PERIODICALS, may 
find a booklet published by the Biological Council of the U.K. (obtainable from the Institute of Biology, 
4 1 Queen's Gate, London, S. W.7, England, U.K.) useful, since it sets out the WORLD LIST abbreviations for 
most biological journals with notes of the USASI abbreviations where these differ. CHEMICAL ABSTRACTS 
publishes quarterly supplements of additional abbreviations. The following points of reference style for 
THE BIOLOGICAL BULLETIN differ from USASI (or modified WORLD LIST) usage: 

A. Journal abbreviations, and book titles, all underlined (for italics) 

B. All components of abbreviations with initial capitals (not as European usage in WORLD LIST e.g. 
J. Cell. Comp. Physiol. NOT/ cell. comp. Physiol.) 

C. All abbreviated components must be followed by a period, whole word components must not (i.e. 
J. Cancer Res.) 

D. Space between all components (e.g. J. Cell. Comp. Physiol., not J.Cell.Comp.Physiol.) 

E. Unusual words in journal titles should be spelled out in full, rather than employing new abbrevi- 
ations invented by the author. For example, use Rit Visindafjelags Islendinga without abbreviation. 

F. All single word journal titles in full (e.g. Veliger, Ecology, Brain). 

G. The order of abbreviated components should be the same as the word order of the complete title 
(i.e. Proc. and Trans, placed where they appear, not transposed as in some BIOLOGICAL ABSTRACTS 
listings). 

H. A few well-known international journals in their preferred forms rather than WORLD LIST or 
USASI usage (e.g. Nature, Science, Evolution NOT Nature, Lond., Science, N.Y.; Evolution, Lancaster, 
Pa.) 

6. Reprints, charges. Authors will be charged the excess over $100 of the total of (a) $30 for each 
printed page beyond 15, (b) $30 for each table, (c) $15 for each formula more complex than a single line 
with simple subscripts or superscripts, and (d) $ 1 5 for each figure, with figures on a single plate all consid- 
ered one figure and parts of a single figure on separate sheets considered separate figures. Reprints may be 
ordered at time of publication and normally will be delivered about two to three months after the issue 
date. Authors (or delegates or foreign authors) will receive page proofs of articles shortly before publication. 
They will be charged the current cost of printers' time for corrections to these (other than corrections of 
printers' or editors' errors). 



- -** 



THE 



BIOLOGICAL BULLETIN 



PUBLISHED BY 
THE MARINE BIOLOGICAL LABORATORY 



Editorial Board 



RUSSELL F. DOOLITTLE, University of California 

at San Diego 

WILLIAM R. ECKBERG, Howard University 
ROBERT D. GOLDMAN, Northwestern University 

C. K. GOVIND, Scarborough Campus, University 

ofToronto 

JUDITH P. GRASSLE, Marine Biological Laboratory 

MICHAEL J. GREENBERG, C. V. Whitney Marine 
Laboratory, University of Florida 

MAUREEN R. HANSON, Cornell University 
JOHN E. HOBBIE, Marine Biological Laboratory 
LIONEL JAFFE, Marine Biological Laboratory 

HOLGER W. JANNASCH, Woods Hole Oceanographic 

Institution 



WILLIAM R. JEFFERY, University of Texas at Austin 

GEORGE M. LANGFORD, University of 

North Carolina at Chapel Hill 

GEORGE D. PAPPAS, University of Illinois at Chicago 
SIDNEY K. PIERCE, University of Maryland 

HERBERT SCHUEL, State University of New York at 

Buffalo 

VIRGINIA L. SCOFIELD, University of California at 
Los Angeles School of Medicine 

LAWRENCE B. SLOBODKJN, State University of New 

York at Stony Brook 

JOHN D. STRANDBERG, Johns Hopkins University 
DONALD P. WOLF, Oregon Regional Primate Center 
SEYMOUR ZIGMAN, University of Rochester 



Editor: CHARLES B. METZ, University of Miami 



AUGUST, 1987 



Printed and Issued by 
LANCASTER PRESS, Inc. 

PRINCE & LEMON STS. 
LANCASTER, PA. 



111 



Marine Biological Laboratory 
LIBRARY 

SEP 141987 



Woods Hole, Mass. 



The BIOLOGICAL BULLETIN is issued six times a year at the 
Lancaster Press, Inc., Prince and Lemon Streets, Lancaster, Penn- 
sylvania. 

Subscriptions and similar matter should be addressed to The 
Biological Bulletin, Marine Biological Laboratory, Woods Hole, 
Massachusetts. Single numbers, $20.00. Subscription per volume 
(three issues), $50.00 ($100.00 per year for six issues). 

Communications relative to manuscripts should be sent to Dr. 
Charles B. Metz, Editor, or Pamela Clapp, Assistant Editor, Marine 
Biological Laboratory, Woods Hole, Massachusetts 02543. 



THE BIOLOGICAL BULLETIN (ISSN 0006-3185) 

POSTMASTER: Send address changes to THE BIOLOGICAL BULLETIN, 

Marine Biological Laboratory, Woods Hole, MA 02543. 
Second-class postage paid at Woods Hole, MA, and additional mailing offices. 



LANCASTER PRESS, INC., LANCASTER, PA 



IV 



THE MARINE BIOLOGICAL LABORATORY 
EIGHTY-NINTH REPORT, FOR THE YEAR 1 986 NINETY-NINTH YEAR 

I. TRUSTEES AND STANDING COMMITTEES 1 

II. MEMBERS OF THE CORPORATION 6 

1 . LIFE MEMBERS 6 

2. REGULAR MEMBERS 8 

3. ASSOCIATE MEMBERS 28 

III. CERTIFICATE OF ORGANIZATION 32 

IV. ARTICLES OF AMENDMENT 33 

V. BYLAWS 34 

VI. REPORT OF THE DIRECTOR 39 

VII. REPORT OF THE TREASURER 43 

VIII. REPORT OF THE LIBRARIAN 55 

IX. EDUCATIONAL PROGRAMS 55 

1 . SUMMER 55 

2. SPRING 64 

3. SHORT COURSES 65 

X. RESEARCH AND TRAINING PROGRAMS 67 

1 . SUMMER 67 

2. YEAR-ROUND 76 

XI. HONORS 82 

XII. INSTITUTIONS REPRESENTED 85 

XIII. LABORATORY SUPPORT STAFF 89 



I. TRUSTEES 

Including Action of the 1986 Annual Meeting 
OFFICERS 

PROSSER GIFFORD, Chairman of the Board of Trustees, Woodrow Wilson International Center 

for Scholars, Smithsonian Building, Washington, DC 20560 
DENIS M. ROBINSON, Honorary Chairman of the Board of Trustees, 200 Ocean Lane, Key 

Biscay ne,FL 33 149 

ROBERT MANZ, Treasurer, 1 Spafford Road, Milton, MA 02186 
PAUL R. GROSS, President of the Corporation, Marine Biological Laboratory, Woods Hole, 

MA 02543 
J. RICHARD WHITTAKER, Director of the Laboratory, Marine Biological Laboratory, Woods 

Hole, MA 02543 
DAVID D. POTTER, Clerk, Harvard Medical School, Cambridge, MA 02138 



Copyright 1987, by the Marine Biological Laboratory 

Library of Congress Card No. A38-5 1 8 

(ISSN 0006-3 185) 



2 MARINE BIOLOGICAL LABORATORY 

EMERITI 

JOHN B. BUCK, National Institutes of Health 

AURIN CHASE, Princeton University 

GEORGE H. A. CLOWES, JR., The Cancer Research Institute 

SEYMOUR S COHEN, Woods Hole, Massachusetts 

ARTHUR L. COLWIN, University of Miami 

LAURA HUNTER COLWIN, University of Miami 

D. EUGENE COPELAND, Marine Biological Laboratory 

SEARS CROWELL, Indiana University 

ALEXANDER T. DAIGNAULT, Boston, Massachusetts 

TERU HAYASHI, Miami, Florida 

HOPE HIBBARD, Oberlin College 

LEWIS KLEINHOLZ, Reed College 

MAURICE KRAHL, Tucson, Arizona 

CHARLES B. METZ, University of Miami 

KEITH PORTER, University of Maryland 

C. LADD PROSSER, University of Illinois 
JOHN S. RANKIN, Ashford, Connecticut 
MERYL ROSE, Waquoit, Massachusetts 
JOHN SAUNDERS, JR., SUNY, Albany 
GEORGE T. SCOTT, Woods Hole, Massachusetts 
MARY SEARS, Woods Hole, Massachusetts 
HOMER P. SMITH, Woods Hole, Massachusetts 
CARL C. SPEIDEL, University of Virginia (no mailings) 

ALBERT SZENT-GYORGYI, Marine Biological Laboratory (deceased 10/22/86) 
W. RANDOLPH TAYLOR, University of Michigan 
GEORGE WALD, Woods Hole, Massachusetts 

CLASS OF 1990 

JOHN E. DOWLING, Harvard University 

GERALD FISCHBACH, Washington University School of Medicine 

ROBERT D. GOLDMAN, Northwestern University 

JOHN E. HOBBIE, Marine Biological Laboratory 

RICHARD KENDALL, Massachusetts Governor's Office 

JOAN V. RUDERMAN, Duke University 

ANN E. STUART, University of North Carolina 

D. THOMAS TRIGG, Wellesley, Massachusetts 

CLASS OF 1989 

GARLAND E. ALLEN, Washington University 
PETER B. ARMSTRONG, University of California, Davis 
ROBERT W. ASHTON, Gaston Snow Beekman and Bogue 
JELLE ATEMA, Marine Biological Laboratory 
HARLYN O. HALVORSON, Brandeis University 
JOHN G. HILDEBRAND, University of Arizona 
THOMAS J. HYNES, JR., Meredith and Grew, Inc. 
ROBERT MAINER, The Boston Company 
BIRGIT ROSE, University of Miami 

CLASS OF 1988 

CLAY M. ARMSTRONG, University of Pennsylvania 

JOEL P. DAVIS, Seapuit, Inc. 

ELLEN R. GRASS, The Grass Foundation 



TRUSTEES AND STANDING COMMITTEES 

JUDITH GRASSLE, Marine Biological Laboratory 

HOLGER W. JANNASCH, Woods Hole Oceanographic Institution 

GEORGE M. LANGFORD, University of North Carolina 

ANDREW SZENT-GYORGYI, Brandeis University 

KENSAL VAN HOLDE, Oregon State University 

RICHARD W. YOUNG, Wellesley Hills, Massachusetts 

CLASS OF 1987 

EDWARD A. ADELBERG, Yale University 
JAMES M. CLARK, Shearson/ American Express 
HAROLD GAINER, National Institutes of Health 
WILLIAM T. GOLDEN, New York, New York 
HANS KORNBERG, University of Cambridge 
LASZLO LORAND, Northwestern University 
CAROL L. REINISCH, Tufts University 
HOWARD A. SCHNEIDERMAN, Monsanto Company 
SHELDON J. SEGAL, The Rockefeller Foundation 



STANDING COMMITTEES 

EXECUTIVE COMMITTEE OF THE BOARD OF TRUSTEES 

PROSSER GIFFORD* JUDITH GRASSLE, 1 988 

PAUL R. GROSS* HARLYN O. HALVORSON, 1989 

J. RICHARD WHITTAKER* JOHN G. HILDEBRAND, 1989 

ROBERT MANZ* ANDREW SZENT-GYORGYI, 1988 

JOHN E. DOWLING, 1990 KENSAL VAN HOLDE, 1988 
GERALD FISCHBACH, 1990 

ANIMAL CARE COMMITTEE 

CAROL L. REINISCH, Chairman ROXANNA SMOLOWITZ 

DANIEL ALKON RAYMOND E. STEPHENS 

EDWARD JASKUN J. RICHARD WHITTAKER 

BUILDINGS AND GROUNDS COMMITTEE 

KENYON S. TWEEDELL, Chairman DONALD B. LEHY* 

LAWRENCE B. COHEN THOMAS H. MEEDEL 

RICHARD D. CUTLER* PHILIP PERSON 

ALAN FEIN LIONEL I. REBHUN 

DANIEL L. GILBERT THOMAS S. REESE 

CIFFORD V. HARDING, JR. EVELYN SPIEGEL 
FERENC I. HAROSI 

CAPITAL DEVELOPMENT COMMITTEE 

RICHARD W. YOUNG, Chairman WILLIAM T. GOLDEN 

PROSSER GIFFORD* HARLYN O. HALVORSON 

EMPLOYEE RELATIONS COMMITTEE 

JOHN V. K. HELFRICH, Chairman EDWARD ENOS 

JUDITH ASHMORE WILLIAM A. EVANS 

FLORENCE DWAYNE JOHN B. MACLEOD 



4 MARINE BIOLOGICAL LABORATORY 

FELLOWSHIPS COMMITTEE 

THORU PEDERSON, Chairman EDUARDO MACAGNO 

JUDITH GRASSLE CAROL L. REINISCH 

JOAN E. HOWARD* J. RICHARD WHITTAKER* 
GEORGE M. LANGFORD 

FINANCIAL POLICY AND PLANNING COMMITTEE 
GEORGE H. A. CLOWES, JR., Chairman ROBERT MAINER 

ROBER i W. ASHTON W. NICHOLAS THORNDIKE 

DAVID L. CURRIER* J. RICHARD WHITTAKER 

THOMAS J. HYNES, JR. 

HOUSING, FOOD SERVICE AND DAY CARE COMMITTEE 

JELLE ATEMA, Chairman LouANN KING* 

ROBERT B. BARLOW, JR. THOMAS S. REESE 

GAIL D. BURD JOAN RUDERMAN 

RONALD L. CALABRESE BRIAN M. SALZBERG 

STEPHEN M. HIGHSTEIN SUSAN SZUTS 

INSTITUTIONAL BIOSAFETY 

RAYMOND E. STEPHENS, Chairman DONALD B. LEHY* 

PAUL J. DE WEER JOSEPH MARTYNA 

PAUL T. ENGLUND ANDREW H. MATTOX* 

HARLYN O. HALVORSON* AL SENFT 
PAUL LEE 

INSTRUCTION COMMITTEE 

JUDITH GRASSLE, Chairman* HANS LAUFER 

BRIAN FRY JOAN V. RUDERMAN 

HARLYN O. HALVORSON* BRIAN M. SALZBERG 

JOHN G. HILDEBRAND* ROGER D. SLOBODA 

JOAN E. HOWARD* ANDREW SZENT-GYORGYI* 

INVESTMENT COMMITTEE 

D. THOMAS TRIGG, Chairman ROBERT MANZ* 

PROSSER GIFFORD* JOHN W. SPEER* 

WILLIAM T. GOLDEN W. NICHOLAS THORNDIKE 

MAURICE LAZARUS J. RICHARD WHITTAKER* 

LIBRARY JOINT MANAGEMENT COMMITTEE 

J. RICHARD WHITTAKER, Chairman* JOHN W. SPEER* 

GARLAND E. ALLEN JOHN H. STEELE 

GEORGE D. GRICE 

LIBRARY JOINT USERS COMMITTEE 

GARLAND E. ALLEN, Chairman LAURENCE P. MADIN 

WILFRED B. BRYAN JOHN SCHLEE 

A. FARMANFARMAIAN FREDERIC SERCHUK 

JANE FESSENDEN* OLIVER C. ZAFIRIOU 
LIONEL F. JAFFE 



TRUSTEES AND STANDING COMMITTEES 

MARINE RESOURCES COMMITTEE 

ROBERT D. GOLDMAN, Chairman GEORGE D. PAPPAS 

WILLIAM D. COHEN ROGER D. SLOBODA 

RICHARD D. CUTLER* MELVIN SPIEGEL 

Louis LEIBOVITZ ANTOINETTE STEINACHER 

TOSHIO NARAHASHI JOHN VALOIS* 

RADIATION SAFETY COMMITTEE 

PAUL J. DE WEER, Chairman ANDREW H. MATTOX* 

RICHARD L. CHAPPELL HARRIS RIPPS 

SHERWIN J. COOPERSTEIN RAYMOND E. STEPHENS 

DANIEL S. GROSCH WALTER S. VINCENT 

RESEARCH SERVICES COMMITTEE 

BIRGIT ROSE, Chairman RAYMOND J. LASEK 

ROBERT B. BARLOW, JR. BRYAN D. NOE 

RICHARD D. CUTLER* BRUCE J. PETERSON 

ROBERT D. GOLDMAN JOEL L. ROSENBAUM 

JOHN G. HILDEBRAND RAYMOND E. STEPHENS 

JOAN E. HOWARD* SIDNEY L. TAMM 
SAMUEL S. KOIDE 

RESEARCH SPACE COMMITTEE 

JOSEPH SANGER, Chairman LASZLO LORAND 

CLAY M. ARMSTRONG EDUARDO MACAGNO 

ROBERT D. GOLDMAN JERRY A. MELILLO 

JOAN E. HOWARD* ROGER D. SLOBODA 

DAVID LANDOWNE EVELYN SPIEGEL 

HANS LAUFER STEVEN N. TREISTMAN 

RODOLFO R. LLINAS IVAN VALIELA 

SAFETY COMMITTEE 

JOHN E. HOBBIE, Chairman ALAN M. KUZIRIAN 

DANIEL L. ALKON DONALD B. LEHY* 

D. EUGENE COPELAND ANDREW H. MATTOX* 

RICHARD D. CUTLER* EDWARD A. SADOWSKI 

EDWARD ENOS RAYMOND E. STEPHENS 

ALAN FEIN PAUL A. STEUDLER 
LOUIS M. KERR 

TRUSTEES' COMMITTEES 

AUDIT COMMITTEE 

ROBERT MAINER, Chairman D. THOMAS TRIGG 

ROBERT MANZ* KENSAL VAN HOLDE 

SHELDON J. SEGAL RICHARD W. YOUNG 

INVESTMENT COMMITTEE 

D. THOMAS TRIGG, Chairman ROBERT MANZ* 

WILLIAM T. GOLDEN W. NICHOLAS THORNDIKE 

MAURICE LAZARUS 



6 MARINE BIOLOGICAL LABORATORY 

COMPENSATION COMMITTEE 

GEORGE H. A. CLOWES, JR., Chairman HARLYN O. HALVORSON 

JAMES M. CLARK THOMAS J. HYNES, JR. 

COMMITTEE ON LABORATORY GOALS 

GERALD FISCHBACH, Chairman JOHN E. HOBBIE 

MICHAEL V. L. BENNETT DAVID D. POTTER 

HARLYN O. HALVORSON JOAN V. RUDERMAN 

JOHN G. HILDEBRAND J. RICHARD WHITTAKER* 

CENTRAL CENTENNIAL COMMITTEE 

JAMES D. EBERT, Chairman JOHN PFEIFFER 

PAMELA CLAPP, Assistant KEITH R. PORTER 

GARLAND E. ALLEN C. LADD PROSSER 

ROBERT B. BARLOW JOHN REED 

RICHARD KENDALL D. THOMAS TRIGG 

II. MEMBERS OF THE CORPORATION 

Including Action of the 1986 Annual Meeting 
LIFE MEMBERS 

ABBOTT, MARIE, c/o Katherine Y. Hutchinson, Bunker Hill Road, Andover, CT 06232 
ADOLPH, EDWARD F., University of Rochester, School of Medicine and Dentistry, Rochester, 

NY 14642 

BEAMS, HAROLD W., Department of Zoology, University of Iowa, Iowa City, IA 53342 
BEHRE, ELLINOR, Black Mountain, NC 2871 1 
BERNHEIMER, ALAN W., New York University, College of Medicine, Charlottesville, VA 

22908 
BERTHOLF, LLOYD M., Westminster Village #2114, 2025 E. Lincoln St., Bloomington, IL 

61701 
BISHOP, DAVID W., Department of Physiology, Medical College of Ohio, C. S. 10008, Toledo, 

OH 43699 

BOLD, HAROLD C., Department of Botany, University of Texas, Austin, TX 78712 
BRIDGMAN, A. JOSEPHINE, 7 1 5 Kirk Rd., Decatur, GA 30030 
BUCK, JOHN B., NIH, Laboratory of Physical Biology, Room 1 12, Building 6, Bethesda, MD 

20892 

BURBANCK, MADELINE P., Box 1 5 1 34, Atlanta, GA 30333 
BURBANCK, WILLIAM D., Box 15134, Atlanta, GA 30333 
CARPENTER, RUSSELL L., 60-H Lake St., Winchester, MA 01890 

CHASE, AURIN, Professor of Biology Emeritus, Princeton University, Princeton, NJ 08540 
CLARKE, GEORGE L., Address Unknown 
CLOWES, GEORGE H. A., JR., The Cancer Research Institute, 194 Pilgrim Rd., Boston, MA 

02215 

COHEN, SEYMOUR S., 10 Carrot Hill Rd., Woods Hole, MA 02543 
COLWIN, ARTHUR, 320 Woodcrest Rd., Key Biscayne, FL 33149 
COLWIN, LAURA HUNTER, 320 Woodcrest Rd., Key Biscayne, FL 33149 
COPELAND, D. E., 41 Fern Lane, Woods Hole, MA 02543 
COSTELLO, HELEN M., Carolina Meadows, Villa 1 37, Chapel Hill, NC 275 14 
CROUSE, HELEN, Institute of Molecular Biophysics, Florida State University, Tallahassee, FL 

32306 

* ex-officio 



MEMBERS OF THE CORPORATION 7 

DILLER, IRENE C, Rydal Park, Apartment 660, Rydal, PA 19046 

DILLER, WILLIAM F., Rydal Park, Apartment 660, Rydal, PA 19046 (deceased 2/8/86) 

ELLIOTT, ALFRED M., 428 Lely Palm Ext., Naples, FL 33962-8903 

FAILLA, PATRICIA M., 2149 Loblolly Lane, Johns Island, SC 29455 

FERGUSON, JAMES K. W., 56 Clarkehaven St., Thornhill, Ontario, Canada L4J 2B4 

FISHER, J. MANERY, Department of Biochemistry, University of Toronto, Toronto, Ontario, 

Canada M5S 1 A8 (deceased 9/9/86) 
FRIES, ERIK F. B., 4 1 High Street, Woods Hole, MA 02543 
OILMAN, LAUREN C., Department of Biology, University of Miami, PO Box 24918, Coral 

Gables, FL 33 134 

GREEN, JAMES W., 409 Grand Ave., Highland Park, NJ 08904 

HAMBURGER, VIKTOR, Professor Emeritus, Washington University, St. Louis, MO 63 1 30 
HAMILTON, HOWARD L., Department of Biology, University of Virginia, Charlottesville, VA 

22901 

HIBBARD, HOPE, c/o Jeanne Stephens, 374 Morgan St., Oberlin, OH 44074 
HISAW, F. L., 5925 SW Plymouth Drive, Corvallis, OR 97330 
HOLLAENDER, ALEXANDER, Council for Research Planning, 1717 Massachusetts Ave., NW, 

Washington, DC 20036 

HUMES, ARTHUR, Marine Biological Laboratory, Woods Hole, MA 02543 
JOHNSON, FRANK H., Department of Biology, Princeton University, Princeton, NJ 08540 
KAAN, HELEN W., Royal Megansett Nursing Home, Room 205, PO Box 408, N. Falmouth, 

MA 02556 

KARUSH, FRED, 183 Summit Lane, Bala-Cynwyd, PA 19004 
KILLE, FRANK R., 1 1 1 1 S. Lakemont Ave. #444, Winter Park, FL 32792 
KINGSBURY, JOHN M., Department of Botany, Cornell University, Ithaca, NY 14853 
KLEINHOLZ, LEWIS, Department of Biology, Reed College, Portland, OR 97202 
LAUFFER, MAX A., Department of Biophysics, University of Pittsburgh, Pittsburgh, PA 15260 
LEFEVRE, PAUL G., 1 5 Agassiz Road, Woods Hole, MA 02543 
LEVINE, RACHMIEL, 2024 Canyon Rd., Arcadia, CA 91006 
LOCHHEAD, JOHN H., 49 Woodlawn Rd., London SW 6 6PS, England, U. K. 
LYNN, W. GARDNER, Department of Biology, Catholic University of America, Washington, 

DC 200 17 

MAGRUDER, SAMUEL R., 270 Cedar Lane, Paducah, KY 42001 
MANWELL, REGINALD D., Syracuse University, Lyman Hall, Syracuse, NY 13210 
MARSLAND, DOUGLAS, Broadmead N 1 2, 1 380 1 York Rd., Cockeysville, MD 2 1 030 (deceased 

8/17/86) 

MILLER, JAMES A., 307 Shorewood Drive, E. Falmouth, MA 02536 
MILNE, LORUS J., Department of Zoology, University of New Hampshire, Durham, NH 

03824 

MOORE, JOHN A., Department of Biology, University of California, Riverside, CA 92521 
MOUL, E. T., 43 F. R. Lillie Rd., Woods Hole, MA 02543 
NACE, PAUL F., 5 Bowditch Road, Woods Hole, MA 02543 
PAGE, IRVING H., Box 516, Hyannisport, MA 02647 
POLLISTER, A. W., 313 Broad Street, Harleysville, PA 19438 
PROSSER, C. LADD, Department of Physiology and Biophysics, Burrill Hall 524, University of 

Illinois, Urbana, IL 6 1801 

PROVASOLI, LUIGI, Haskins Laboratories, 165 Prospect Street, New Haven, CT 065 10 
PRYTZ, MARGARET MCDONALD, 21 McCouns Lane, Oyster Bay, NY 1 1771 
RANKIN, JOHN S., JR., Box 97, Ashford, CT 06278 
RENN, CHARLES E., Route 2, Hempstead, MD 21074 
RICHARDS, A. GLENN, 942 Cromwell Ave., St. Paul, MN 55 1 14 
RICHARDS, OSCAR W., Pacific University, Forest Grove, OR 97462 
RONKIN, RAPHAEL R., 3212 McKinley St., NW, Washington, DC 20015 
SCHARRER, BERTA, Department of Anatomy, Albert Einstein College of Medicine, 1 300 Mor- 
ris Park Avenue, Bronx, NY 10461 



8 MARINE BIOLOGICAL LABORATORY 

SCHLESINGER, R. WALTER, University of Medicine and Dentistry of New Jersey, Department 
of Microbiology, Rutgers Medical School, PO Box 101, Piscataway, NJ 08854 

SCHMITT, F. O., Room 16-512, Massachusetts Institute of Technology, Cambridge, MA 02 1 39 

SCOTT, ALLAN C., 1 Nudd St., Waterville, ME 04901 

SCOTT, GEORGE T., 10 Orchard St., Woods Hole, MA 02543 

SHEMIN, DAVID, Department of Biochemistry and Molecular Biology, Northwestern Univer- 
sity, Evanston, IL 60201 

SMITH, HOMER P., 8 Quissett Ave., Woods Hole, MA 02543 

SONNENBLICK, B. P., Department of Zoology and Physiology, Rutgers University, 195 Univer- 
sity Ave., Newark, NJ 07 102 

SPEIDEL, CARL C., 1873 Field Rd., Charlottesville, VA 22903 (no mailings) 

STEINHARDT, JACINTO, 1 508 Spruce St., Berkeley, CA 94709 

STUNKARD, HORACE W., American Museum of Natural History, Central Park West at 79th 
St., New York, NY 10024 

TAYLOR, W. RANDOLPH, Department of Biology, University of Michigan, Ann Arbor, MI 
48109 

TAYLOR, W. ROWLAND, 152 Cedar Park Road, Annapolis, MD 21401 

TEWINKEL, Lois E., 4 Sanderson Ave., Northampton, MA 01060 

TRACER, WILLIAM, The Rockefeller University, 1230 York Ave., New York, NY 10021 

WAINIO, WALTER W., 331 State Road, Princeton, NJ 08540 

WALD, GEORGE, 67 Gardner Road, Woods Hole, MA 02543 

WEISS, PAUL A., Address Unknown 

WICHTERMAN, RALPH, 3 1 Buzzards Bay Ave., Woods Hole, MA 02543 

WIERCINSKI, FLOYD J., Department of Biology, Northwestern Illinois University, Chicago, IL 
60625 

WILBER, CHARLES G., Department of Zoology, Colorado State University, Fort Collins, CO 
80523 

YOUNG, D. B., 1 1 37 Main St., N. Hanover, MA 02357 

ZINN, DONALD J., PO Box 589, Falmouth, MA 02541 

ZORZOLI, ANITA, 18 Wilbur Blvd., Poughkeepsie, NY 12603 

ZWEIFACH, BENJAMIN W., c/o Ames, University of California, La Jolla, CA 92037 

REGULAR MEMBERS 

ACHE, BARRY W., Whitney Marine Laboratory, University of Florida, Rt. 1, Box 121, St. 
Augustine, FL 32086 

ACHESON, GEORGE H., 25 Quissett Ave., Woods Hole, MA 02543 

ADAMS, JAMES A., Department of Biological Sciences, Tennessee State University 3500 John 
Merritt Blvd., Nashville, TN 37203 

ADELBERG, EDWARD A., Department of Human Genetics, Yale University Medical School, 
PO Box 3333, New Haven, CT 065 10 

AFZELIUS, BJORN, Wenner-Gren Institute, University of Stockholm, Stockholm, Sweden 

ALBERTE, RANDALL S., University of Chicago, Barnes Laboratory, 5630 S. Ingleside Ave., 
Chicago, IL 60637 

ALKON, DANIEL, Section on Neural Systems, Laboratory of Biophysics, NIH, Marine Biologi- 
cal Laboratory, Woods Hole, MA 02543 

ALLEN, GARLAND E., Department of Biology, Washington University, St. Louis, MO 63130 

ALLEN, NINA S., Department of Biology, Wake Forest University, Box 7325, Reynolds Sta- 
tion, Winston-Salem, NC 27109 

ALLEN, ROBERT D., Department of Biology, Dartmouth College, Hanover, NH 03755 (de- 
ceased 3/23/86) 

AMATNIEK, ERNEST, 4797 Boston Post Rd., Pelham Manor, NY 10803 

ANDERSON, EVERETT, Department of Anatomy, LHRBB, Harvard Medical School, Boston, 
MA 02115 

ANDERSON, J. M., 1 10 Roat St., Ithaca, NY 14850 



MEMBERS OF THE CORPORATION 

ARMET-KiBEL, CHRISTINE, Biology Department, University of Massachusetts-Boston, Bos- 
ton, MA 02 125 

ARMSTRONG, CLAY M., Department of Physiology, Medical School, University of Pennsylva- 
nia, Philadelphia, PA 19174 

ARMSTRONG, PETER B., Department of Zoology, University of California, Davis, CA 95616 

ARNOLD, JOHN M., Pacific Biomedical Research Center, 209 Snyder Hall, 2538 The Mall 
Honolulu, HI 96822 

ARNOLD, WILLIAM A., 102 Balsam Rd., Oak Ridge, TN 37830 

ASHTON, ROBERT W., Gaston Snow Beekman and Bogue, 14 Wall St., New York, NY 10005 

ATEMA, JELLE, Marine Biological Laboratory, Woods Hole, MA 02543 

ATWOOD, KIMBALL C, PO Box 673, Woods Hole, MA 02543 

AUGUSTINE, GEORGE JR., Section of Neurobiology, Department of Biological Sciences, Uni- 
versity of Southern California, Los Angeles, CA 90089-037 1 

AUSTIN, MARY L., 506'/2 N. Indiana Ave., Bloomington, IN 47401 

AYERS, DONALD E., Marine Biological Laboratory, Woods Hole, MA 02543 

BACON, ROBERT, PO Box 723, Woods Hole, MA 02543 

BAKER, ROBERT G., New York University Medical Center, 550 First Ave., New York, NY 
10016 

BALDWIN, THOMAS O., Department of Biochemistry and Biophysics, Texas A&M University, 
College Station, TX 77843 

BANG, BETSY, 76 F. R. Lillie Rd., Woods Hole, MA 02543 

BARKER, JEFFERY L., National Institutes of Health, Bldg. 36, Room 2002, Bethesda, MD 
20892 

BARLOW, ROBERT B., JR., Institute for Sensory Research, Syracuse University, Merrill Lane, 
Syracuse, NY 13210 

BARRY, DANIEL T., Department of Physical Medicine and Rehabilitation, ID204, University 
of Michigan Hospital, Ann Arbor, MI 48109-0042 

BARRY, SUSAN R., Department of Physical Medicine and Rehabilitation, ID204, University 
of Michigan Hospital, Ann Arbor, MI 48109-0042 

BARTELL, CLELMER K., 2000 Lake Shore Drive, New Orleans, LA 70122 

BARTH, LUCENA J., 26 Quissett Ave., Woods Hole, MA 02543 (deceased 7/26/86) 

BARTLETT, JAMES H., Department of Physics, Box 1921, University of Alabama, Tuscaloosa, 
AL 35489 

BASS, ANDREW H., Seely Mudd Hall, Department of Neurobiology and Behavior, Cornell 
University, Ithaca, NY 14853 

BATTELLE, BARBARA-ANNE, Whitney Marine Laboratory, Rt. 1, Box 121, St. Augustine, FL 
32086 

BAUER, G. ERIC, Department of Anatomy, University of Minnesota, Minneapolis, MN 55455 

BEAUGE, Luis ALBERTO, Institute de Investigacion Medica, Casilla de Correo 389, 5000 Cor- 
doba, Argentina 

BECK, L. V., School of Experimental Medicine, Department of Pharmacology, Indiana Uni- 
versity, Bloomington, IN 47401 

BEGENISICH, TED, Department of Physiology, University of Rochester, Rochester NY 14642 

BEGG, DAVID A., LHRRB, Harvard Medical School, 45 Shattuck St., Boston, MA 02 1 1 5 

BELL, EUGENE, Department of Biology, Massachusetts Institute of Technology, 77 Massachu- 
setts Ave., Cambridge, MA 02 1 39 

BENJAMIN, THOMAS L., Department of Pathology, Harvard Medical School, 25 Shattuck St., 
Boston, MA 021 15 

BENNETT, M. V. L., Albert Einstein College of Medicine, Department of Neuroscience, 1300 
Morris Park Ave., Bronx, NY 10461 

BENNETT, MIRIAM F., Department of Biology, Colby College, Waterville, ME 04901 

BERG, CARL J., JR., Marine Biological Laboratory, Woods Hole, MA 02543 

BERNE, ROBERT M., University of Virginia, School of Medicine, Charlottesville, VA 22908 

BEZANILLA, FRANCISCO, Department of Physiology, University of California, Los Angeles, 
CA 90052 

BIGGERS, JOHN D., Department of Physiology, Harvard Medical School, Boston, MA 021 15 



10 MARINE BIOLOGICAL LABORATORY 

BISHOP, STEPHEN H., Department of Zoology, Iowa State University, Ames, IA 50010 

BLAUSTEIN, MORDECAI P., Department of Physiology, School of Medicine, University of 
Maryland, 655 W. Baltimore Street, Baltimore, MD 21201 

BLOOM, KERRY S., Department of Biology, University of North Carolina, Chapel Hill, NC 
27514 

BODIAN, DAVID, Address Unknown 

BODZNICK, DAVID A., Department of Biology, Wesleyan University, Middletown, CT 06457 

BOETTIGER. EDWARD G., 29 Juniper Point, Woods Hole, MA 02543 

BOGORAD, LAWRENCE, The Biological Laboratories, Harvard University, Cambridge, MA 
02 1 38 (resigned 8/8/86) 

BOOLOOTIAN, RICHARD A., Science Software Systems, Inc., 3576 Woodcliff Rd., Sherman 
Oaks, CA 9 1403 

BOREI, HANS G., Long Cove, Stanley Point Road, Minturn, ME 04659 

BORGESE, THOMAS A., Department of Biology, Lehman College, CUNY, Bronx, NY 10468 

BORISY, GARY G., Laboratory of Molecular Biology, University of Wisconsin, Madison, WI 
53715 

BOSCH, HERMAN F., PO Box 542, Woods Hole, MA 02543 

BOTKIN, DANIEL, Department of Biology, University of California, Santa Barbara, CA 93106 
(resigned 3/86) 

BOWLES, FRANCIS P., PO Box 674, Woods Hole, MA 02543 

BOYER, BARBARA C, Department of Biology, Union College, Schenectady, NY 12308 

BRANDHORST, BRUCE P., Biology Department, McGill University, 1205 Ave. Dr. Penfield, 
Montreal, P. Q., Canada H3A 1 B 1 

BREHM, PAUL, Department of Physiology, Tufts Medical School, Boston, MA 021 1 1 

BRINLEY, F. J., Neurological Disorders Program, NINCDS, 716 Federal Building, Bethesda, 
MD 20892 

BROWN, JOEL E., Department of Ophthalmology, Box 8096 Sciences Center, Washington Uni- 
versity, 660 S. Euclid Ave., St. Louis, MO 63 1 10 

BROWN, STEPHEN C., Department of Biological Sciences, SUNY, Albany, NY 12222 

BURD, GAIL DEERIN, Department of Molecular and Cellular Biology, Biosciences West, 
Room 305, University of Arizona, Tucson, AZ 85721 

BURDICK, CAROLYN J., Department of Biology, Brooklyn College, Brooklyn, NY 11210 

BURGER, MAX, Department of Biochemistry, Biocenter, Klingelbergstrasse 70, CH-4056 Ba- 
sel, Switzerland 

BURKY, ALBERT, Department of Biology, University of Dayton, Dayton, OH 45469 

BURSTYN, HAROLD LEWIS, 216 Bradford Parkway, Syracuse, NY 13224 

BURSZTAJN, SHERRY, Neurology Department Program in Neuroscience, Baylor College of 
Medicine, Houston, TX 77030 

BUSH, LOUISE, 7 Snapper Lane, Falmouth, MA 02540 

CALABRESE, RONALD L., Department of Biology, Emory University, 1555 Pierce Drive, At- 
lanta, GA 30322 

CANDELAS, GRACIELA C., Department of Biology, University of Puerto Rico, Rio Piedras, PR 
00931 

CAREW, THOMAS J., Department of Psychology, Yale University, PO Box 1 1 A, Yale Station, 
New Haven, CT 06520 

CARIELLO, Lucio, Stazione Zoologica, Villa Comunale, Naples, Italy 

CARLSON, FRANCIS D., Department of Biophysics, Johns Hopkins University, Baltimore, MD 
21218 

CASE, JAMES, Department of Biological Sciences, University of California, Santa Barbara, CA 
93106 

CASSIDY, REV. J. D., St. Rose Priory, Springfield, KY 40069 

CEBRA, JOHN J., Department of Biology, Leidy Labs, G-6, University of Pennsylvania, Phila- 
delphia, PA 19174 

CHAET, ALFRED B., University of West Florida, Pensacola, FL 32504 

CHAMBERS, EDWARD L., Department of Physiology and Biophysics, University of Miami, 
School of Medicine, PO Box 016430, Miami, FL 33101 



MEMBERS OF THE CORPORATION 1 1 

CHANG, DONALD C., Department of Physiology and Molecular Biophysics, Baylor College of 

Medicine, One Baylor Plaza, Houston, TX 77030 
CHAPPELL, RICHARD L., Department of Biological Sciences, Hunter College Box 210, 695 

Park Ave., New York, NY 10021 

CHAUNCEY, HOWARD H., 30 Falmouth St., Wellesley Hills, MA 02 1 8 1 
CHARLTON, MILTON P., Physiology Department MSB, University of Toronto, Toronto, On- 
tario, Canada M5S 1 A8 

CHILD, FRANK M., Department of Biology, Trinity College, Hartford, CT 06 106 
CHISHOLM, REX L., Dept. of Cell Biology and Anatomy, Northwestern University Medical 

School, 303 E. Chicago Avenue, Chicago, IL 6061 1 
CITKOWITZ, ELENA, 410 Livingston St., New Haven, CT 065 1 1 
CLARK, A. M., 48 Wilson Rd., Woods Hole, MA 02543 

CLARK, ELOISE E. Vice President for Academic Affairs, Bowling Green State University, Bowl- 
ing Green, OH 43403 

CLARK, HAYS, Property Management Ltd., 125 Mason St., Greenwich, CT 06830 
CLARK, JAMES M., Shearson Lehman Brothers Inc., Two World Trade Center, 105th Floor, 

New York, NY 10048 

CLARK, WALLIS H., JR., Bodega Marine Lab, PO Box 247, Bodega Bay, CA 94923 
CLAUDE, PHILIPPA, Primate Center, Capitol Court, Madison, WI 53706 
CLAY, JOHN R., Marine Biological Laboratory, Woods Hole, MA 02543 
CLOWES, GEORGE H. A., JR., The Cancer Research Institute, 194 Pilgrim Rd., Boston, MA 

02215 
CLUTTER, MARY, Senior Science Advisor, Office of the Director, Room 5 1 8, National Science 

Foundation, Washington, DC 20550 

COBB, JEWELL P., President, California State University, Fullerton, CA 92634 
COHEN, ADOLPH L, Department of Ophthalmology, School of Medicine, Washington Univer- 
sity, 660 S. Euclid Ave., St. Louis, MO 631 10 
COHEN, CAROLYN, Rosenstiel Basic Medical Sciences Research Center, Brandeis University, 

Waltham, MA02154 
COHEN, LAWRENCE B., Department of Physiology, Yale University School of Medicine, B- 

106 SHM, PO Box 3333, New Haven, CT 065 10-8026 
COHEN, MAYNARD, Department of Neurological Sciences, Rush Medical College 600 South 

Paulina, Chicago, IL 606 1 2 
COHEN, ROCHELLE S., Department of Anatomy, University of Illinois at Chicago, 808 S. 

Wood Street, Chicago, I L 606 1 2 
COHEN, WILLIAM D., Department of Biological Sciences, Hunter College, 695 Park Ave., New 

York, NY 10021 

COLE, JONATHAN J., Institute for Ecosystems Studies, Cary Arboretum, Millbrook, NY 12545 
COLEMAN, ANNETTE W., Division of Biology and Medicine, Brown University, Providence, 

RI02912 

COLLIER, JACK R., Department of Biology, Brooklyn College, Brooklyn, NY 11210 
COLLIER, MARJORIE McCANN, Biology Department, Saint Peter's College, Kennedy Boule- 
vard, Jersey City, NJ 07306 
COOK, JOSEPH A., The Edna McConnell Clark Foundation, 250 Park Ave., New York, NY 

10017 
COOPERSTEIN, S. J., University of Connecticut, School of Medicine, Farmington Ave., Far- 

mington, CT 06032 

CORLISS, JOHN O., Department of Zoology, University of Maryland, College Park, MD 20742 
CORNELL, NEAL W., 6428 Bannockburn Drive, Bethesda, MD 208 1 7 
CORNMAN, IVOR, 10A Orchard St., Woods Hole, MA 02543 (resigned 12/4/86) 
CORNWALL, MELVIN C., JR., Department of Physiology L714, Boston University School of 

Medicine, 80 E. Concord St., Boston, MA 02 1 1 8 

CORSON, DAVID WESLEY, JR., 1034 Plantation Lane, Mt. Pleasant, SC 29464 
CORWIN, JEFFREY T., Bekesy Lab of Neurobiology, 1993 East- West Road, University of Ha- 
waii, Honolulu, HI 96822 
COSTELLO, WALTER J., College of Medicine, Ohio University, Athens, OH 45701 



12 MARINE BIOLOGICAL LABORATORY 

COUCH, ERNEST F., Department of Biology, Texas Christian University, Fort Worth, TX 
76129 

CREMER-BARTELS, GERTRUD, Universitats Augenklinik, 44 Munster, West Germany 

CROW, TERRY J., Department of Physiology, University of Pittsburgh, School of Medicine, 
Pittsburgh, PA 15261 

CROWELL, SEARS, Department of Biology, Indiana University, Bloomington, IN 47405 

CROWTHER, ROBERT, Marine Biological Laboratory, Woods Hole, MA 02543 

CURRIER, DAVID L., PO Box 2476, Vineyard Haven, MA 02568 

DAIGNAULT, ALEXANDER T., 280 Beacon St., Boston, MA 021 16 

DAN, KATSUMA, Tokyo Metropolitan Union, Meguro-ku, Tokyo, Japan 

D'AVANZO, CHARLENE, School of Natural Science, Hampshire College, Amherst, MA 01002 

DAVID, JOHN R., Seeley G. Mudd Building, Room 504, Harvard Medical School, 250 Long- 
wood Ave., Boston, MA 02 1 1 5 

DAVIDSON, ERIC H., Division of Biology, California Institute of Technology, Pasadena, CA 
91125 

DAVIS, BERNARD D., 23 Clairemont Road, Belmont, MA 02 1 78 

DAVIS, JOEL P., Seapuit, Inc., PO Box G, Osterville, MA 02655 

DAW, NIGEL W., 78 Aberdeen Place, Clayton, MO 63105 

DEGROOF, ROBERT C, RR#1 Box 343, Green Lane, PA 18054 

DEHAAN, ROBERT L., Department of Anatomy, Emory University, Atlanta, GA 30322 

DELANNEY, Louis E., Institute for Medical Research, 2260 Clove Drive, San Jose, CA 95 128 

DEPHILLIPS, HENRY A., JR., Department of Chemistry, Trinity College, Hartford, CT 06 106 

DETERRA, NOEL, 2 1 5 East 1 5th St., New York, NY 1 0003 

DETTBARN, WOLF-DIETRICH, Department of Pharmacology, School of Medicine, Vanderbilt 
University, Nashville, TN 37 127 

DE WEER, PAUL J., Department of Physiology, School of Medicine, Washington University, 
St. Louis, MO 63 110 

DISCHE, ZACH ARIAS, Eye Institute, College of Physicians and Surgeons, Columbia University, 
639 W. 165 St., New York, NY 10032 (dropped 3/86) 

DIXON, KEITH E., School of Biological Sciences, Flinders University, Bedford Park, South 
Australia 

DONELSON, JOHN E., Department of Biochemistry, University of Iowa, Iowa City IA 52242 

DOWDALL, MICHAEL J., Department of Zoology, School of Biological Sciences, University of 
Nottingham, University Park, Nottingham N672 UH, England, U. K. 

DOWLING, JOHN E., The Biological Laboratories, Harvard University, 16 Divinity St., Cam- 
bridge, MA 02 1 38 

DuBois, ARTHUR BROOKS, John B. Pierce Foundation Laboratory, 290 Congress Ave., New 
Haven, CT 065 19 

DUDLEY, PATRICIA L., Department of Biological Sciences, Barnard College, Columbia Uni- 
versity, New York, NY 10027 

DUNCAN, THOMAS K., Department of Environmental Science, Nichols College, Dudley, MA 
01570 

DUNHAM, PHILIP B., Department of Biology, Syracuse University, Syracuse, NY 13210 

DUNLAP, KATHLEEN, Department of Psychology, Tufts Medical School, Boston, MA 021 1 1 

EBERT, JAMES D., Office of the President, Carnegie Institute of Washington 1530 P St., NW, 
Washington, DC 20008 

ECKBERG, WILLIAM R., Department of Zoology, Howard University, Washington, DC 20059 

ECKERT, ROGER O., Department of Zoology, University of California, Los Angeles, CA 90024 
(deceased 6/1 8/86) 

EDDS, KENNETH T., Department of Anatomical Sciences, SUNY, Buffalo, NY 14214 

EDER, HOWARD A., Albert Einstein College of Medicine, 1300 Morris Park Ave., Bronx, NY 
10461 

EDWARDS, CHARLES, NIAADK/NIH, Rm. 403, Bldg. 10, Bethesda, MD 20892 

EGYUD, LASZLO G., 1 8 Skyview, Newton, MA 02 1 50 

EHRENSTEIN, GERALD, NIH, Bethesda, MD 20892 



MEMBERS OF THE CORPORATION 13 

EHRLICH, BARBARA E., Department of Physiology, Albert Einstein College of Medicine, 1 300 
Morris Park Ave., Bronx, NY 10461 

EISEN, ARTHUR Z., Chief of Division of Dermatology, Washington University, St. Louis, MO 
63110 

EISENMAN, GEORGE, Department of Physiology, University of California Medical School, Los 
Angeles, CA 90024 

ELDER, HUGH YOUNG, Institute of Physiology, University of Glasgow, Glasgow, Scotland, 
U.K. 

ELLIOTT, GERALD F., The Open University Research Unit, Foxcombe Hall, Berkeley Rd., 
Boars Hill. Oxford, England, U. K. 

ENGLUND, PAUL T., Department of Biological Chemistry, Johns Hopkins School of Medicine, 
Baltimore, MD 2 1205 

EPEL, DAVID, Hopkins Marine Station, Pacific Grove, CA 93950 

EPSTEIN, HERMAN T., Department of Biology, Brandeis University, Waltham, MA 02254 

ERULKAR, SOLOMON D., 318 Kent Rd., Bala Cynwyd, PA 19004 

ESSNER, EDWARD S., Kresege Eye Institute, Wayne State University, 540 E. Canfield Ave., 
Detroit, MI 48201 

FARMANFARMAIAN, A., Department of Biological Sciences, Nelson Biological Laboratory, 
Rutgers University, PO Box 1059, Piscataway, NJ 08854 

FEIN, ALAN, Laboratory of Sensory Physiology, Marine Biological Laboratory, Woods Hole, 
MA 02543 

FEINMAN, RICHARD D., Box 8, Department of Biochemistry, SUNY Health Science Center, 
Brooklyn, NY 11203 

FELDMAN, SUSAN C., Department of Anatomy, University of Medicine and Dentistry of New 
Jersey, New Jersey Medical School, 100 Bergen St., Newark, NJ 07103 

FERGUSON, F. P., National Institute of General Medical Science, NIH, Bethesda, MD 20892 

FESSENDEN, JANE, Marine Biological Laboratory, Woods Hole, MA 02543 

FESTOFF, BARRY W., Neurology Service ( 1 27), Veterans Administration Medical Center, 480 1 
Linwood Blvd., Kansas City, MO 64128 

FINKELSTEIN, ALAN, Albert Einstein College of Medicine, 1 300 Morris Park Ave., Bronx, NY 
10461 

FISCHBACH, GERALD, Department of Anatomy and Neurobiology, Washington University 
School of Medicine, St. Louis, MO 631 10 

FISCHMAN, DONALD A., Department of Cell Biology and Anatomy, Cornell University Medi- 
cal College, 1 300 York Ave., New York, NY 1002 1 

FISHMAN, HARVEY M., Department of Physiology, University of Texas Medical Branch, Gal- 
veston,TX 77550 

FLANAGAN, DENNIS, 12 Gay St., New York, NY 10014 

Fox, MAURICE S., Department of Biology, Massachusetts Institute of Technology, Cambridge, 
MA 02 138 

FRANK, PETER W., Department of Biology, University of Oregon, Eugene, OR 97403 

FRANZINI, CLARA, Department of Biology G-5, School of Medicine, University of Pennsylva- 
nia, Philadelphia, PA 19174 

FRAZIER, DONALD T., Department of Physiology and Biophysics, University of Kentucky 
Medical Center, Lexington, KY 40536 

FREEMAN, ALAN R., Department of Physiology, Temple University, 3420 N. Broad St., Phila- 
delphia, PA 19140 (resigned 3/86) 

FREEMAN, GARY L., Department of Zoology, University of Texas, Austin, TX 78 172 

FREINKEL, NORBERT, Center for Endocrinology, Metabolism & Nutrition, Northwestern Uni- 
versity Medical School, 303 E. Chicago Avenue, Chicago, IL 6061 1 

FRENCH, ROBERT J., Department of Medical Physiology, University of Calgary, 3330 Hospital 
Dr., NW, Calgary, Alberta T2N 4N1 Canada 

FREYGANG, WALTER J., JR., 6247 29th St., NW, Washington, DC 20015 

FRY, BRIAN, Marine Biological Laboratory, Woods Hole, MA 02543 

FUKUI, YOSHIO, Department of Cell Biology and Anatomy, Northwestern University Medical 
School, Chicago, IL 60201 



14 MARINE BIOLOGICAL LABORATORY 

FULTON, CHANDLER M, Department of Biology, Brandeis University, Waltham, MA 02 1 54 
FURSHPAN, EDWIN J., Department of Neurophysiology, Harvard Medical School, Boston, MA 

02115 
FUSELER, JOHN W., Department of Biology, University of Southwestern Louisiana, Lafayette, 

LA 70504 

FUTRELLE, ROBERT P., College of Computer Science, Northeastern University, 360 Hunting- 
ton Avenue, Boston, MA 02 1 1 5 
FYE, PAUL, PO Box 309, Woods Hole, MA 02543 

GABRIEL, MORDECAI, Department of Biology, Brooklyn College, Brooklyn, NY 11210 
GADSBY, DAVID C, Laboratory of Cardiac Physiology, The Rockefeller University, 1 230 York 

Avenue, New York, NY 1002 1 
GAINER, HAROLD, Section of Functional Neurochemistry, NIH, Bldg. 36 Room 2A21, 

Bethesda, MD 20892 

GALATZER-LEVY, ROBERT M., 180 N. Michigan Avenue, Chicago, IL 60601 
GALL, JOSEPH G., Carnegie Institution, 1 1 5 West University Parkway, Baltimore, MD 21210 
GALLANT, PAUL E., Laboratory of Preclinical Studies, Bldg. 36, NIAAA/NIH, 1250 Washing- 
ton Ave., Rockville, MD 20892 
GASCOYNE, PETER, Department of Experimental Pathology, Box 85E, University of Texas 

System Cancer Center, M. D. Anderson Hospital and Tumor Institute, Texas Medical 

Center, 6723 Bertner Avenue, Houston, TX 77030 
GELFANT, SEYMOUR, Department of Dermatology, Medical College of Georgia, Augusta, GA 

30904 

GELPERIN, ALAN, Department of Biology, Princeton University, Princeton, NJ 08540 
GERMAN, JAMES L., Ill, The New York Blood Center, 310 East 67th St., New York, NY 1002 1 
GIBBS, MARTIN, Institute for Photobiology of Cells and Organelles, Brandeis University, Wal- 
tham, MA 02 154 

GIBLIN, ANNE E., Ecosystems Center, Marine Biological Laboratory, Woods Hole, MA 02543 
GIBSON, A. JANE, Wing Hall, Cornell University, Ithaca, NY 14850 
GIFFORD, PROSSER. The Wilson Center, Smithsonian Building, 1000 Jefferson Drive, SW, 

Washington, DC 20590 
GILBERT, DANIEL L., NIH, Laboratory of Biophysics, NINCDS, Bldg. 36, Room 2A-29, 

Bethesda, MD 20892 

GIUDICE, GIOVANNI, Via Archirafi 22, Palermo, Italy 
GLUSMAN, MURRAY, Department of Psychiatry, Columbia University, 722 W. 1 68th St., New 

York, NY 10032 

GOLDEN, WILLIAM T., 40 Wall St., New York, NY 10005 
GOLDMAN, DAVID E., 63 Loop Rd., Falmouth, MA 02540 
GOLDMAN, ROBERT D., Department of Cell Biology and Anatomy, Northwestern University, 

303 E. Chicago Ave., Chicago, IL 6061 1 

GOLDSMITH, PAUL K. 55 1 1 Oakmont Avenue, Bethesda, MD 20034 
GOLDSMITH, TIMOTHY H., Department of Biology, Yale University, New Haven, CT 065 10 
GOLDSTEIN, MOISE H., JR., EE & CS Department, Johns Hopkins University, Baltimore, MD 

21218 
GOODMAN, LESLEY JEAN, Department of Biological Sciences, Queen Mary College, Mile End 

Road, London, El 4NS, England, U. K. 

GOUDSMIT, ESTHER M., Department of Biology, Oakland University, Rochester, MI 48063 
GOULD, ROBERT MICHAEL, Institute for Basic Research in Developmental Disabilities, 1050 

Forest Hill Rd., Staten Island, NY 10314 
GOULD, STEPHEN J., Museum of Comparative Zoology, Harvard University, Cambridge, MA 

02138 
GOVIND, C. K., Zoology Department-Scarborough, University of Toronto, 1265 Military 

Trail, West Hill, Ontario, Canada, MIC 1A4 
GRAF, WERNER, Rockefeller University, New York, NY 10021 
GRAHAM, HERBERT, 36 Wilson Rd., Woods Hole, MA 02543 
GRANT, PHILIP, Department of Biology, University of Oregon, Eugene, OR 97403 
GRASS, ALBERT, The Grass Foundation, 77 Reservoir Rd., Quincy, MA 02 1 70 



MEMBERS OF THE CORPORATION 15 

GRASS, ELLEN R., The Grass Foundation, 77 Reservoir Rd., Quincy, MA 02170 
GRASSLE, JUDITH, Marine Biological Laboratory, Woods Hole, MA 02543 
GREEN, JONATHAN P., Department of Biology, Roosevelt University, 430 S. Michigan Ave- 
nue, Chicago, IL 60605 

GREENBERG, EVERETT PETER, Department of Microbiology, Stocking Hall, Cornell Univer- 
sity, Ithaca, NY 14853 
GREENBERG, MICHAEL J., Whitney Marine Laboratory, Rt. 1, Box 121, St. Augustine, FL 

32086 
GREIF, ROGER L., Department of Physiology, Cornell University, Medical College New York, 

NY 10021 

GRIFFIN, DONALD R., The Rockefeller University, 1230 York Ave., New York, NY 1002 1 
GROSCH, DANIEL S., Department of Genetics, Gardner Hall, North Carolina State University, 

Raleigh, NC 27607 
GROSS, PAUL R., President and Director, Marine Biological Laboratory, Woods Hole, MA 

02543 

GROSSMAN, ALBERT, New York University, Medical School, New York, NY 10016 
GUNNING, A. ROBERT, PO Box 165, Falmouth, MA 02541 
GWILLIAM, G. P., Department of Biology, Reed College, Portland, OR 97202 
HALL, LINDA M., Department of Genetics, Albert Einstein College of Medicine, 1300 Morris 

Park Ave., Bronx, NY 1046 1 
HALL, ZACK W., Department of Physiology, University of California, San Francisco, CA 

94143 

HALVORSON, HARLYN O., Rosenstiel Basic Medical Sciences Research Center, Brandeis Uni- 
versity, Waltham, MA 02 1 54 
HAMLETT, NANCY VIRGINIA, Department of Biology, Swarthmore College, Swarthmore, PA 

19081 
HANNA, ROBERT B., College of Environmental Science and Forestry, SUNY, Syracuse, NY 

13210 
HARDING, CLIFFORD V., JR., Kresege Eye Institute, Wayne State University, 540 E. Canfield, 

Detroit, MI 48201 
HAROSI, FERENC I., Laboratory of Sensory Physiology, Marine Biological Laboratory, Woods 

Hole, MA 02543 

HARRIGAN, JUNE F., 7415 Makaa Place, Honolulu, HI 96825 
HARRINGTON, GLENN W., Department of Microbiology, School of Dentistry, University of 

Missouri, 650 E. 25th St., Kansas City, MO 64108 
HARRIS, ANDREW L., Department of Biophysics, Johns Hopkins University, 34th & Charles 

Sts., Baltimore, MD 2 12 18 
HASCHEMEYER, AUDREY E. V., Department of Biological Sciences, Hunter College, 695 Park 

Ave., New York, NY 10021 

HASTINGS, J. W., The Biological Laboratories, Harvard University, Cambridge, MA 02138 
HAUSCHKA, THEODORE S., RD1, Box 781, Damariscotta, ME 04543 
HAYASHI, TERU, 7105 SW 1 12 Place, Miami, FL 33173 
HAYES, RAYMOND L., JR., Dept. of Anatomy, Howard University, College of Medicine, 520 

W St., NW, Washington, DC 20059 

HENLEY, CATHERINE, 5225 Pooks Hill Rd., #1 127 North, Bethesda, MD 20034 
HEPLER, PETER K., Department of Botany, University of Massachusetts, Amherst, MA 01003 
HERNDON, WALTER R., University of Tennessee, Department of Biology, Knoxville, TN 

37996-1100 

HESSLER, ANITA Y., 5795 Waverly Ave., La Jolla, CA 92037 
HEUSER, JOHN, Department of Biophysics, Washington University, School of Medicine, St. 

Louis, MO 63 110 

HIATT, HOWARD H., Brigham and Women's Hospital, 75 Francis Street, Boston, MA 021 15 
HIGHSTEIN, STEPHEN M., Department of Otolaryngology, Washington University, St. Louis, 

MO63110 
HILDEBRAND, JOHN G., Arizona Research Laboratories, Division of Neurobiology, 603 

Gould-Simpson Science Building, University of Arizona, Tucson, AZ 85721 



16 MARINE BIOLOGICAL LABORATORY 

HILL, SUSAN D., Department of Zoology, Michigan State University, E. Lansing, MI 48824 
HILLIS-COLINVAUX, LLEWELLYA, Department of Zoology, The Ohio State University, 484 W 

1 2th Ave., Columbus, OH 432 1 

HILLMAN, PETER, Department of Biology, Hebrew University, Jerusalem, Israel 
HINEGARDNER, RALPH T., Division of Natural Sciences, University of California Santa Cruz, 

CA 95064 
HINSCH, GERTRUDE, W., Department of Biology, University of South Florida, Tampa, FL 

33620 

HoBBUi, JOHN E., Ecosystems Center, Marine Biological Laboratory, Woods Hole, MA 02543 
HODGE, ALAN J., Marine Biological Laboratory, Woods Hole, MA 02543 
HOFFMAN, JOSEPH, Department of Physiology, School of Medicine, Yale University, New 

Haven, CT 065 10 

HOLLYFIELD, JOE G., Baylor School of Medicine, Texas Medical Center, Houston, TX 77030 
HOLTZMAN, ERIC, Department of Biological Sciences, Columbia University, New York, NY 

10017 

HOLZ, GEORGE G., JR., Department of Microbiology, SUNY, Syracuse, NY 13210 
HOSKIN, FRANCIS C. G., Department of Biology, Illinois Institute of Technology, Chicago, IL 

60616 
HOUGHTON, RICHARD A., Ill, Ecosystems Center, Marine Biological Laboratory, Woods 

Hole, MA 02543 

HOUSTON, HOWARD E., 2500 Virginia Ave., NW, Washington, DC 20037 
HOWARD, JOAN E., Marine Biological Laboratory, Woods Hole, MA 02543 
HOWARTH, ROBERT, Section of Ecology & Systematics, Corson Hall, Cornell University, Ith- 
aca, NY 14853 
HOY, RONALD R., Section of Neurobiology and Behavior, Cornell University, Ithaca, NY 

14850 

HUBBARD, RUTH, 67 Gardner Road, Woods Hole, MA 02543 
HUFNAGEL, LINDA A., Department of Microbiology, University of Rhode Island, Kingston, 

RI 02881 

HUMMON, WILLIAM D., Department of Zoology, Ohio University, Athens, OH 45701 
HUMPHREYS, SUSIE H., Kraft Research and Development, 801 Waukegan Rd., Glenview, IL 

60025 

HUMPHREYS, TOM D., University of Hawaii, PBRC, 41 Ahui St., Honolulu, HI 968 13 
HUNTER, BRUCE W., Box 32 1 , Lincoln Center, MA 1 773 
HUNTER, ROBERT D., Department of Biological Sciences, Oakland University, Rochester, NY 

48063 

HUNZIKER, HERBERT E., Esq., PO Box 547, Falmouth, MA 0254 1 
HURWITZ, CHARLES, Basic Science Research Lab, Veterans Administration Hospital, Albany, 

NY 12208 
HURWITZ, JERARD, Memorial Sloan Kettering Institute, 1275 York Avenue, New York NY 

11021 
HUXLEY, HUGH E., Medical Research Council, Laboratory of Molecular Biology, Cambridge, 

England, U. K. 

HYNES, THOMAS J., JR., Meredith and Grew, Inc., 125 High Street, Boston, MA 02 110 
ILAN, JOSEPH, Department of Anatomy, Case Western Reserve University, Cleveland, OH 

44106 
INGOGLIA, NICHOLAS, Department of Physiology, New Jersey Medical School, 100 Bergen St., 

Newark, NJ 07 103 

INOUE, SADUYKI, McGill University Cancer Centre, Department of Anatomy, 3640 Univer- 
sity St., Montreal, PQ, Canada, H3A 2B2 

INOUE, SHINYA, Marine Biological Laboratory, Woods Hole, MA 02543 
ISSADORIDES, MARIETTA R., Department of Psychiatry, University of Athens, Monis Petraki 

8, Athens, 140 Greece 

ISSELBACHER, KURT J., Massachusetts General Hospital, 32 Fruit Street, Boston, MA 02 1 1 4 
IZZARD, COLIN S., Department of Biological Sciences, SUNY, Albany, NY 12222 
JACOBSON, ANTONE G., Department of Zoology, University of Texas, Austin, TX 78712 



MEMBERS OF THE CORPORATION 17 

JAFFE, LIONEL, Marine Biological Laboratory, Woods Hole, MA 02543 

JAHAN-PARWAR, BEHRUS, Center for Laboratories & Research, New York State Department 

of Health, Empire State Plaza, Albany, NY 12201 

JANNASCH, HOLGER W., Woods Hole Oceanographic Institution, Woods Hole, MA 02543 
JEFFERY, WILLIAM R., Department of Zoology, University of Texas, Austin, TX 78712 
JENNER, CHARLES E., Department of Zoology, University of North Carolina, Chapel Hill, NC 

27514 

JONES, MEREDITH L., Division of Worms, Museum of Natural History, Smithsonian Institu- 
tion, Washington, DC 20560 
JOSEPHSON, ROBERT K., School of Biological Sciences, University of California, Irvine, CA 

92664 
KABAT, E. A., Department of Microbiology, College of Physicians and Surgeons Columbia 

University, 630 West 168th St., New York, NY 10032 

KALEY, GABOR, Department of Physiology, Basic Sciences Building, New York Medical Col- 
lege, Valhalla, NY 10595 
KALTENBACH, JANE, Department of Biological Sciences, Mount Holyoke College, South Had- 

ley,MA01075 
KAMINER, BENJAMIN, Department of Physiology, School of Medicine, Boston University, 80 

East Concord St., Boston, MA 02 1 1 8 

KAMMMER, ANN E., Department of Zoology, Arizona State University, Tempe, AZ 85281 
KANE, ROBERT E., University of Hawaii, PBRC, 41 Ahui St., Honolulu, HI 96813 
KANESHIRO, EDNA S., Department of Biological Sciences, University of Cincinnati, Cincin- 
nati, OH 45221 

KAO, CHIEN-YUAN, Department of Pharmacology (Box 29), SUNY, Downstate Medical Cen- 
ter, 450 Clarkson Avenue, Brooklyn, NY 1 1203 

KAPLAN, EHUD, The Rockefeller University, 1230 York Ave., New York, NY 10021 
KARAKASHIAN, STEPHEN J., Apt. 16-F, 165 West 9 1st St., New York, NY 10024 
KARLIN, ARTHUR, Department of Biochemistry and Neurology, Columbia University, 630 

West 168th St., New York, NY 10032 
KATZ, GEORGE M., Fundamental and Experimental Research, Merck, Sharpe and Dohme 

Rahway, NJ 07065 
KEAN, EDWARD L., Department of Ophthalmology and Biochemistry, Case Western Reserve 

University, Cleveland, OH 44101 
KELLEY, DARCY BRISBANE, Department of Biological Sciences, 1018 Fairchild, Columbia 

University, New York, NY 10032 
KELLY, ROBERT E., Department of Anatomy, College of Medicine, University of Illinois, PO 

Box 6998, Chicago, IL 60680 

KEMP, NORMAN E., Department of Zoology, University of Michigan, Ann Arbor, MI 48104 
KENDALL, JOHN P., Faneuil Hall Associates, One Boston Place, Boston, MA 02108 
KENDALL, RICHARD, 26 Green Harbor Rd., East Falmouth, MA 02536 
KEYNAN, ALEXANDER, Hebrew University, Jerusalem, Israel 

KJEHART, DANIEL P., Department of Cellular and Developmental Biology, Harvard Univer- 
sity, 16 Divinity Avenue, Cambridge, MA 02138 

KLEIN, MORTON, Department of Microbiology, Temple University, Philadelphia, PA 19103 
KLOTZ, I. M., Department of Chemistry, Northwestern University, Evanston, IL 60201 
KOIDE, SAMUEL S., Population Council, The Rockefeller University, 66th St. and York Ave., 

New York, NY 10021 
KONIGSBERG, IRWIN R., Department of Biology, Gilmer Hall, University of Virginia, Char- 

lottesville, VA 22903 
KORNBERG, SIR HANS, Department of Biochemistry, University of Cambridge, Tennis Court 

Rd., Cambridge, CB2 7QW, England, U. K. 
KOSOWER, EDWARD M., Ramat-Aviv, Tel Aviv, 69978 Israel 
KRAHL, M. E., 2783 W. Casas Circle, Tucson, AZ 8574 1 
KRANE, STEPHEN M., Massachusetts General Hospital, Boston, MA 02 1 14 
KRASSNER, STUART M., Department of Developmental and Cell Biology, University of Cali- 
fornia, Irvine, CA 927 1 7 



18 MARINE BIOLOGICAL LABORATORY 

KRAUSS, ROBERT, FASEB, 9650 Rockville Pike, Bethesda, MD 20814 

KRAVITZ, EDWARD A., Department of Neurobiology, Harvard Medical School, 25 Shattuck 

St., Boston, MA 02 115 
KRIEBEL, MAHLON E., Department of Physiology, B.S.B., Upstate Medical Center, 766 Irving 

A ve., Syracuse, NY 13210 

KRIEG, WENDELL J. S., 1236 Hinman, Evanston, IL 60602 (resigned 3/86) 
KRISTAN, WILLIAM B., JR., Department of Biology B-022, University of California San Diego, 

San Diego, CA 92093 
KUHNS, WILLIAM J., University of North Carolina, 512 Faculty Lab Office, Bldg. 231-H, 

Chapel Hill, NC 275 14 

KUSANO, KJYOSHI, Illinois Institute of Technology, Department of Biology, 3300 South Fed- 
eral St., Chicago, IL 606 1 6 

KUZIRIAN, ALAN M., Laboratory of Biophysics, NINCDS-NIH, Marine Biological Labora- 
tory, Woods Hole, MA 02543 

LADERMAN, AIMLEE, PO Box 689, Woods Hole, MA 02543 

LAMARCHE, PAUL H., Eastern Maine Medical Center, 489 State St., Bangor, ME 04401 
LANDIS, DENNIS M. D., Department of Developmental Genetics and Anatomy, Case Western 

Reserve Medical School, 2119 Abington Road, Cleveland, OH 44106 
LANDIS, STORY C., Department of Pharmacology, Case Western Reserve University Medical 

School, 21 19 Abington Road, Cleveland, OH 44106 

LANDOWNE, DAVID, Department of Physiology, Yale University School of Medicine, 333 Ce- 
dar St., New Haven, CT 065 10 
LANGFORD, GEORGE M., Department of Physiology, Medical Sciences Research Wing 206H, 

University of North Carolina, Chapel Hill, NC 275 14 
LASER, RAYMOND J., Case Western Reserve University, Department of Anatomy, Cleveland, 

OH 44 106 

LASTER, LEONARD, University of Oregon, Health Sciences Center, Portland, OR 97201 
LAUFER, HANS, Biological Sciences Group U-42, University of Connecticut, Storrs, CT 06268 
LAZAROW, PAUL B., The Rockefeller University, 1 230 York Avenue, New York, NY 1002 1 
LAZARUS, MAURICE, Federated Department Stores, Inc., 50 Cornhill, Boston, MA 02108 
LEADBETTER, EDWARD R., Department of Molecular and Cell Biology, U-131, University of 

Connecticut, Storrs, CT 06268 
LEDERBERG, JOSHUA, President, The Rockefeller University, 1230 York Ave., New York, NY 

10021 
LEDERHENDLER, IZJA I., Laboratory of Biophysics, Marine Biological Laboratory, Woods 

Hole, MA 02543 
LEE, JOHN J., Department of Biology, City College of CUNY, Convent Ave. and 138th St., 

New York, NY 10031 

LEHY, DONALD B., Marine Biological Laboratory, Woods Hole, MA 02543 
LEIBOVITZ, Louis, Laboratory for Marine Animal Health, Marine Biological Laboratory, 

Woods Hole, MA 02543 

LEIGHTON, JOSEPH, 1201 Waverly Rd., Gladwyne, PA 19035 
LEIGHTON, STEPHEN, NIH, Bldg. 13 3W13, Bethesda, MD 20892 
LEINWAHN, LESLIE ANN, Department of Microbiology and Immunology, 1300 Morris Park 

Ave., Bronx, NY 10461 
LERMAN, SIDNEY, Laboratory for Ophthalmic Research, Emory University, Atlanta, GA 

30322 

LERNER, AARON B., Yale University, School of Medicine, New Haven, CT 065 10 
LESTER, HENRY A., 156-29 California Institute of Technology, Pasadena, CA 91 125 
LEVIN, JACK, Clinical Pathology Service, VA Hospital- 1 1 3A, 4 1 50 Clement St., San Francisco, 

CA 94121 
LEVINTHAL, CYRUS, Department of Biological Sciences, Columbia University, 435 Riverside 

Drive, New York, NY 10025 
LEVITAN, HERBERT, Department of Zoology, University of Maryland, College Park, MD 

20742 



MEMBERS OF THE CORPORATION 19 

LINCK, RICHARD W., Department of Anatomy, Jackson Hall, University of Minnesota, 321 

Church Street, S.E., Minneapolis, MN 55455 

LING, GILBERT, 307 Berkeley Road, Merion, PA 19066 (dropped 9/1/86) 
LIPICKY, RAYMOND J., Department of Cardio-Renal/HFD 1 10, FDA Bureau of Drugs, Rm. 

16B-45, 5600 Fishers Lane, Rockville, MD 20857 

LISMAN, JOHN E., Department of Biology, Brandeis University, Waltham, MA 02 1 54 
Liuzzi, ANTHONY, Department of Physics, University of Lowell, Lowell, MA 01854 
LLINAS, RODOLFO R., Department of Physiology and Biophysics, New York University Medi- 
cal Center, 550 First Ave., New York, NY 10016 
LOEWENSTEIN, WERNER R., Department of Physiology and Biophysics, University of Miami, 

PO Box 1 6430, Miami, FL 33 1 1 
LOEWUS, FRANK A., Institute of Biological Chemistry, Washington State University, Pullman, 

WA99164 
LOFTFIELD, ROBERT B., Department of Biochemistry, School of Medicine, University of New 

Mexico, 900 Stanford, NE, Albuquerque, NM 87 1 3 1 

LONDON, IRVING M., Massachusetts Institute of Technology, Cambridge, MA 02139 
LONGO, FRANK J., Department of Anatomy, University of Iowa, Iowa City, IA 52442 
LORAND, LASZLO, Department of Biochemistry and Molecular Biology, Northwestern Uni- 
versity, Evanston, IL 60201 

LUCKENBILL-EDDS, LOUISE, 1 55 Columbia Ave., Athens, OH 4570 1 
LURIA, SALVADOR E., 48 Peacock Farm Rd., Lexington, MA 02173 
MACAGNO, EDUARDO R., 1003B Fairchild, Columbia University, New York, NY 10022 
MACNiCHOL, E. F., JR., 45 Brewster Street, Cambridge, MA 02 1 38 
MAGLOTT-DUFFIELD, DONNA R. S., 1014 Baltimore Road, Rockville, MD 20851 
MAIENSCHEIN, JANE ANN, Department of Philosophy, Arizona State University, Tempe, AZ 

85281 

MAINER, ROBERT, The Boston Company, One Boston Place, Boston, MA 02 108 
MALBON, CRAIG CURTIS, Department of Pharmacological Sciences, Health Sciences Center, 

SUNY, Stony Brook, Stony Brook, NY 1 1794-865 1 

MALKIEL, SAUL, Allergic Diseases, Inc., 130 Lincoln St., Worcester, MA 01605 
MANALIS, RICHARDS., Department of Biological Sciences, Purdue University, 2101 Coliseum 

Blvd., East, Ft. Wayne, IN 46805 

MANGUM, CHARLOTTE P., Department of Biology, College of William and Mary, Williams- 
burg, VA 23 185 
MARGULIS, LYNN, Department of Biology, Boston University, 2 Cummington St., Boston, 

MA 022 15 

MARINUCCI, ANDREW C., 26 Woodlawn Ave., North Brunswick, NJ 08902 
MARSH, JULIAN B., Department of Biochemistry and Physiology, Medical College of Pennsyl- 
vania, 3300 Henry Ave., Philadelphia, PA 19129 

MARTIN, LOWELL V., Marine Biological Laboratory, Woods Hole, MA 02543 
MARTINEX-PALOMO, ADOLFO, Seccion de Patologia Experimental, Cinvesav-ipn, 17000 

Mexico, D. F. A. P., 14-740, Mexico 
MASER, MORTON, PO Box EM, Woods Hole, MA 02543 

MASTROIANNI, LUIGI, JR., Department of Obstetrics and Gynecology, University of Pennsyl- 
vania, Philadelphia, PA 19174 
MATHEWS, RITA W., Department of Medicine, New York University Medical Center, 550 

First Ave., New York, NY 10016 
MATTESON, DONALD R., Department of Physiology, G4, School of Medicine, University of 

Pennsylvania, Philadelphia, PA 19104 
MAUTNER, HENRY G., Department of Biochemistry and Pharmacology, Tufts University, 1 36 

Harrison Ave., Boston, MA 021 1 1 

MAUZERALL, DAVID, The Rockefeller University, 1230 York Ave., New York, NY 10021 
MAZIA, DANIEL, Hopkins Marine Station, Pacific Grove, CA 93950 

MAZZELLA, LUCIA, Laboratorio di Ecologia del Benthos, Stazione Zoologica di Napoli, P.ta 
S. Pietro 80077, Ischia Porto (NA), Italy 



20 MARINE BIOLOGICAL LABORATORY 

McCANN, FRANCES, Department of Physiology, Dartmouth Medical School, Hanover, NH 

03755 
McCLOSKEY, LAWRENCE R., Department of Biology, Walla Walla College, College Place, WA 

99324 

MCLAUGHLIN, JANE A., PO Box 187, Woods Hole, MA 02543 
McMAHON, ROBERT F., Department of Biology, Box 19498, University of Texas, Arlington, 

TX76019 
MEEDEL, THOMAS, Boston University Marine Program, Marine Biological Laboratory, 

Woods Hole, MA 02543 

MEINERTZHAGEN, IAN A. Department of Psychology, Life Sciences Center, Dalhousie Univer- 
sity, Halifax, Nova Scotia B3H 45 1 , Canada 

MEINKOTH, NORMAN A., 43 1W Woodland Avenue, Springfield, PA 19064 
MEISS, DENNIS E., 462 Solano Avenue, Hayward, CA 94541 
MELILLO, JERRY A., Ecosystems Center, Marine Biological Laboratory, Woods Hole, MA 

02543 

MELLON, RICHARD P., PO Box 187, Laughlintown, PA 15655 
MELLON, DEFOREST, JR., Department of Biology, University of Virginia, Charlottesville, VA 

22903 
MENZEL, RANDOLF, Institut fir Tierphysiologie, Free Universitat of Berlin, 1000 Berlin 41, 

Federal Republic of Germany 

METUZALS, JANIS, Department of Anatomy, Faculty of Medicine, University of Ottawa, Ot- 
tawa, Ontario KIN 9A9, Canada 

METZ, CHARLES B., 7220 SW 124th St., Miami, FL 33156 

MILKMAN, ROGER, Department of Zoology, University of Iowa, Iowa City, IA 52242 
MILLS, ERIC L., Oceanography Dept., Dalhousie University, Halifax, Nova Scotia B3H 4J1, 

Canada 

MILLS, ROBERT, 10315 44th Avenue, W 12 H Street, Bradenton, FL 33507-1535 
MITCHELL, RALPH, Pierce Hall, Harvard University, Cambridge, MA 02138 
MIYAMOTO, DAVID M., Department of Biology, Drew University, Madison NJ 07940 
MIZELL, MERLE, Department of Biology, Tulane University, New Orleans, LA 701 18 
MONROY, ALBERTO, Stazione Zoologica, Villa Comunale, Naples, Italy (deceased 8/23/86) 
MOORE, JOHN W., Department of Physiology, Duke University Medical Center, Durham, NC 

27710 
MOORE, LEE E., Department of Physiology and Biophysics, University of Texas, Medical 

Branch, Galveston, TX 77550 

MORIN, JAMES G., Department of Biology, University of California, Los Angeles, CA 90024 
MORRELL, FRANK, Department of Neurological Sciences, Rush Medical Center, 1753 W. 

Congress Parkway, Chicago, IL 606 1 2 

MORRILL, JOHN B., JR., Division of National Sciences, New College, Sarasota, FL 33580 
MORSE, RICHARD S., 1 93 Winding River Rd., Wellesley, MA 02 1 8 1 
MORSE, ROBERT W., Box 574, N. Falmouth, MA 02556 
MORSE, STEPHEN SCOTT, The Rockefeller University, 1230 York Ave., Box 2, New York, NY 

10021-6399 
MOSCONA, A. A., Department of Biology, University of Chicago, 920 East 58th St., Chicago, 

IL 60637 

MOTE, MICHAEL I., Department of Biology, Temple University, Philadelphia, PA 19122 
MOUNTAIN, ISABEL, Vinson Hall #1 12, 6251 Old Dominion Drive, McLean, VA 22101 
MULLINS, LORIN J., University of Maryland, School of Medicine, Baltimore MD 21201 
MUSACCHIA, XAVIER J., Graduate School, University of Louisville, Louisville, KY 40292 
NABRIT, S. M., 686 Beckwith St., SW, Atlanta, GA 30314 
NADELHOFFER, KNUTE, Marine Biological Laboratory, Woods Hole, MA 02543 
NAKA, KEN-ICHI, National Institute for Basic Biology, Okazaki, Japan 444 
NAKAJIMA, SHIGEHIRO, Department of Biological Sciences, Purdue University, West Lafay- 
ette, IN 47907 

NAKAJIMA, YASUKO, Department of Biological Sciences, Purdue University, West Lafayette, 
IN 47907 



MEMBERS OF THE CORPORATION 21 

NARAHASHI, TOSHIO, Department of Pharmacology, Medical Center, Northwestern Univer- 
sity, 303 East Chicago Ave., Chicago, IL 6061 1 

NASATIR, MAIMON, Department of Biology, University of Toledo, Toledo, OH 43606 

NELSON, LEONARD, Department of Physiology, Medical College of Ohio, Toledo, OH 43699 

NELSON, MARGARET C, 119 Forest Home Drive, Ithaca, NY 14850 

NICHOLLS, JOHN G., Biocenter, Klingelbergstr 70, Basel 4056, Switzerland 

NICOSIA, SANTO V., Department of Pathology, University of South Florida, College of Medi- 
cine, Box 1 1, 12901 North 30th St., Tampa, FL 33612 

NIELSEN, JENNIFER B. K., Merck, Sharp & Dohme Laboratories, Bldg. 50-G, Room 226, Rah- 
way, NJ 07065 

NOE, BRYAN D., Department of Anatomy, Emory University, Atlanta, GA 30345 

OBAID, ANA LIA, Department of Physiology and Pharmacy, University of Pennsylvania, 4001 
Spruce St., Philadelphia, PA 19104 

OCHOA, SEVERO, 530 East 72nd St., New York, NY 10021 

ODUM, EUGENE, Department of Zoology, University of Georgia, Athens, GA 30701 

OERTEL, DONATA, Department of Neurophysiology, University of Wisconsin, 283 Medical 
Science Bldg., Madison, WI 53706 

O'HERRON, JONATHAN, Lazard Freres and Company, 1 Rockefeller Plaza, New York, NY 
10020 

OLINS, ADA L., University of Tennessee-Oak Ridge, Graduate School of Biomedical Sciences, 
Biology Division ORNL, PO Box Y, Oak Ridge, TN 37830 

OLINS, DONALD E., University of Tennessee-Oak Ridge, Graduate School of Biomedical Sci- 
ences, Biology Division ORNL, PO Box Y, Oak Ridge, TN 37830 

O'MELIA, ANNE F., 16 Evergreen Lane, Chappaqua, New York 10514 

OSCHMAN, JAMES L., 9 George Street, Woods Hole, 02543 

PALMER, JOHN D., Department of Zoology, University of Massachusetts, Amherst, MA 01002 

PALTI, YORAM, Department of Physiology and Biophysics, Israel Institute of Technology, 12 
Haaliya St., BAT-GALIM, POB 9649, Haifa, Israel 

PANT, HARISH C., Laboratory of Preclinical Studies, National Institute on Alcohol Abuse and 
Alcoholism, 12501 Washington Ave., Rockville, MD 20852 

PAPPAS, GEORGE D., Department of Anatomy, College of Medicine, University of Illinois, 
808 South Wood St., Chicago, IL 606 1 2 

PARDEE, ARTHUR B., Department of Pharmacology, Harvard Medical School, Boston, MA 
02115 

PARDY, ROSEVELT L., School of Life Sciences, University of Nebraska, Lincoln, NE 68588 

PARMENTIER, JAMES L., Becton Dickinson, PO Box 12016, Research Triangle Park, NC 
27709 

PASSANO, LEONARD M., Department of Zoology, Birge Hall, University of Wisconsin, Madi- 
son, WI 53706 

PEARLMAN, ALAN L., Department of Physiology, School of Medicine, Washington University, 
St. Louis, MO 63 110 

PEDERSON, THORU, Worcester Foundation for Experimental Biology, Shrewsbury, MA 1 545 

PERKINS, C. D., 400 Hilltop Terrace, Alexandria, VA 22301 

PERSON, PHILIP, Oral Health Director, Research Testing Labs, Inc., 167 E. 2nd St., Hunting- 
ton Station, NY 11746 

PETERSON, BRUCE J., 82 Hillcrest Dr., Falmouth, MA 02540 

PETHIG, RONALD, School of Electronic Engineering Science, University College of N. Wales, 
Dean St., Bangor, Gwynedd, LL57 IUT, U. K. 

PETTIBONE, MARIAN H., Division of Worms, W-213, Smithsonian Institution, Washington, 
DC 20560 

PFOHL, RONALD J., Department of Zoology, Miami University, Oxford, OH 45056 

PIERCE, SIDNEY K., JR., Department of Zoology, University of Maryland, College Park, MD 
20740 

POINDEXTER, JEANNE S., Science Division, Long Island University, Brooklyn Campus, 
Brooklyn, NY 11201 

POLLARD, HARVEY B., NIH, F Building 10, Room 10B17, Bethesda, MD 20892 



22 MARINE BIOLOGICAL LABORATORY 

POLLARD, THOMAS D., Department of Cell Biology and Anatomy, Johns Hopkins University, 

725 North Wolfe St., Baltimore, MD 21205 

POLLOCK, LELAND W., Department of Zoology, Drew University, Madison, NJ 07940 
POOLE, ALAN F., 1 14 Metoxit Road, Waquoit, MA 02536 
PORTER, BEVERLY H., 13617 Glenoble Drive, Rockville, MD 20853 
PORTER, KEITH R., 4009 St. John's Lane, Ellicott City, MD 2 1043 
PORTER, MARY E., Department MCD Biology, Campus Box 347, University of Colorado, 

Boulder, CO 80309 

POTTER, DAVID, Department of Neurobiology, Harvard Medical School, Boston, MA 021 15 
POTTS, WILLIAM T., Department of Biology, University of Lancaster, Lancaster, England, 

U.K. 

POUSSART, DENIS, Department of Electrical Engineering, Universite Laval, Quebec, Canada 
PRATT, MELANIE M., Department of Anatomy and Cell Biology, University of Miami School 

of Medicine (R 124), PO Box 016960, Miami, FL 33101 
PRENDERGAST, ROBERT A., Department of Pathology and Ophthalmology, Johns Hopkins 

University, Baltimore, MD 21205 

PRESLEY, PHILLIP H., Carl Zeiss, Inc., 1 Zeiss Drive, Thornwood, NY 10594 
PRICE, CARL A., Waksman Institute of Microbiology, Rutgers University, PO Box 759, Piscat- 

away,NJ 08854 
PRICE, CHRISTOPHER H., Biological Science Center, Boston University, 2 Cummington St., 

Boston, MA 022 1 5 
PRIOR, DAVID J., Department of Biological Sciences, University of Kentucky, Lexington, KY 

40506 

PRUSCH, ROBERT D., Department of Life Sciences, Gonzaga University, Spokane, WA 99258 
PRZYBYLSKI, RONALD J., Case Western Reserve University, Department of Anatomy, Cleve- 
land, OH 44 104 
PURVES, DALE, Department of Anatomy, Washington University School of Medicine, 660 S. 

Euclid Ave., St. Louis, MO 631 10 
QUIGLEY, JAMES, Department of Microbiology and Immunology Box 44, SUNY Downstate 

Medical Center, 450 Clarkson Ave., Brooklyn, NY 1 1203 
RABIN, HARVEY, DuPont Biomed. Prod.-BRL-2, 331 Treble Cove Road, No. Billerica, MA 

01862 

RAFF, RUDOLF A., Department of Biology, Indiana University, Bloomington, IN 47405 
RAKOWSKI, ROBERT F., Department of Physiology and Biophysics, UHS/The Chicago Medi- 
cal School, 3333 Greenbay Rd., N. Chicago, IL 60064 
RAMON, FIDEL, Dept. de Fisiologia y Biofisca, Centrol de Investigacion y de, Estudius Avan- 

zados del Ipn, Apurtado Postal 14-740, Mexico, D. F. 07000 
RANZI, SILVIO, Sez Zoologia Sc Nat, Via Coloria 26, 120 1 33, Milano, Italy 
RATNER, SARAH, Department of Biochemistry, Public Health Research Institute, 455 First 

Ave., New York, NY 10016 
REBHUN, LIONEL I., Department of Biology, Gilmer Hall, University of Virginia, Charlottes- 

ville,VA 22901 
REDDAN, JOHN R., Department of Biological Sciences, Oakland University, Rochester, MI 

48063 

REESE, BARBARA F., Marine Biological Laboratory, Woods Hole, MA 02543 
REESE, THOMAS S., Marine Biological Laboratory, Woods Hole, MA 02543 
REINER, JOHN M., Albany Medical College of Union University, Department of Biochemistry, 

Albany, NY 12208 
REINISCH, CAROL L., Tufts University School of Veterinary Medicine, 203 Harrison Avenue, 

Boston, MA 02 115 
REUBEN, JOHN P., Department of Biochemistry, Merck Sharp and Dohme, PO Box 2000, 

Rahway, NJ 07065 
REYNOLDS, GEORGE T., Department of Physics, Jadwin Hall, Princeton University, 

Princeton, NJ 08540 

RICE, ROBERT V., 30 Burnham, Dr., Falmouth, MA 02540 
RICKLES, FREDERICK R., University of Connecticut, School of Medicine, VA Hospital, New- 

ington,CT06111 



MEMBERS OF THE CORPORATION 23 

RIPPS, HARRIS, Department of Ophthalmology, University of Illinois at Chicago, College of 
Medicine, 1855 W. Taylor Street, Chicago, IL 6061 1 

ROBERTS, JOHN L., Department of Zoology, University of Massachusetts, Amherst, MA 
01002 

ROBINSON, DENIS M., 200 Ocean Lane Drive, Key Biscayne, FL 33149 

ROCKSTEIN, MORRIS, 335 Fluvia Ave., Miami, FL 33 1 34 

ROSBASH, MICHAEL, Rosenstiel Center, Department of Biology, Brandeis University, Wal- 
tham, MA 02 154 

ROSE, BIRGIT, Department of Physiology R-430, University of Miami School of Medicine, PO 
Box 016430, Miami, FL 33149 

ROSE, S. MERYL, Box 309W, Waquoit, MA 02536 

ROSENBAUM, JOEL L., Department of Biology, Kline Biology Tower, Yale University, New 
Haven, CT 06520 

ROSENBERG, PHILIP, School of Pharmacy, Division of Pharmacology, University of Connecti- 
cut, Storrs, CT 06268 

ROSENBLUTH, JACK, Department of Physiology, New York University School of Medicine, 
550 First Ave., New York, NY 10016 

ROSENBLUTH, RAJA, 3380 West 5th Ave., Vancouver 8, British Columbia V6R 1R7, Canada 

ROSLANSKY, JOHN, Box 208, Woods Hole, MA 02543 

ROSLANSKY, PRISCILLA F., Box 208, Woods Hole, MA 02543 

Ross, WILLIAM N., Department of Physiology, New York Medical College, Valhalla, NY 
10595 

ROTH, JAY S., Division of Biological Sciences, Section of Biochemistry and Biophysics, Uni- 
versity of Connecticut, Storrs, CT 06268 

ROWLAND, LEWIS P., Neurological Institute, 710 West 168th St., New York, NY 10032 

RUDERMAN, JOAN V., Department of Zoology, Duke University, Durham, NC 27706 

RUSHFORTH, NORMAN B., Case Western Reserve University, Department of Biology, Cleve- 
land, OH 44 106 

RUSSELL-HUNTER, W. D., Department of Biology, Lyman Hall 029, Syracuse University, 
Syracuse, NY 13210 

SAFFO, MARY BETH, Center for Marine Studies, 273 Applied Sciences, University of Califor- 
nia, Santa Cruz, CA 95064 

SAGER, RUTH, Sidney Farber Cancer Institute, 44 Binney St., Boston, MA 02 1 1 5 

SALAMA, GUY, Department of Physiology, University of Pittsburgh, Pittsburgh, PA 15261 

SALMON, EDWARD D., Department of Zoology, University of North Carolina, Chapel Hill, 
NC 27514 

SALZBERG, BRIAN M., Department of Physiology, University of Pennsylvania, 4010 Locust 
St., Philadelphia, PA 19174 

SANBORN, RICHARD C., 5862 North Olney St., Indianapolis, IN 46220 

SANDERS, HOWARD, Woods Hole Oceanographic Institution, Woods Hole, MA 02543 

SANGER, JEAN M., Department of Anatomy, School of Medicine, University of Pennsylvania, 
36th and Hamilton Walk, Philadelphia, PA 19174 

SANGER, JOSEPH, Department of Anatomy, School of Medicine, University of Pennsylvania, 
36th and Hamilton Walk, Philadelphia, PA 19174 

SATO, EIMEI, Department of Animal Science, Faculty of Agriculture, Kyoto University, Kyoto 
606, Japan 

SATO, HIDEMI, Sugashima Marine Biological Laboratory, Nagoya University, Sugashima-cho, 
Toba-chi, Mie-Ken 517, Japan 

SATTELLE, DAVID B., AFRC Unit-Department of Zoology, University of Cambridge, Down- 
ing St., Cambridge CB2 3EJ, England, U. K. 

SAUNDERS, JOHN, JR., Department of Biological Sciences, SUNY, Albany, NY 12222 

SAZ, ARTHUR K., Medical and Dental Schools, Georgetown University, 3900 Reservoir Rd., 
NW, Washington, DC 2005 1 

SCHACHMAN, HOWARD K., Department of Molecular Biology, University of California, 

Berkeley, CA 94720 

SCHATTEN, GERALD P., Integrated Microscopy Facility for Biomedical Research, University 
of Wisconsin, 1 1 17 W. Johnson St., Madison, WI 53706 



24 MARINE BIOLOGICAL LABORATORY 

SCHATTEN, HEIDI, Department of Zoology, University of Wisconsin, Madison WI 53706 
SCHIFF, JEROME A., Institute for Photobiology of Cells and Organelles, Brandeis University, 

Waltham, MA 02 1 54 
SCHMEER, ARLENE C, Mercene Cancer Research Hospital of Saint Raphael, New Haven, CT 

06511 

SCHNAPP, BRUCE J., Marine Biological Laboratory, Woods Hole, MA 02543 
SCHNEIDER, E. GAYLE, Department of Obstetrics and Gynecology, Yale University School of 

Medicine, 333 Cedar St., New Haven, CT 065 10 
SCHNEIDERMAN, HOWARD A., Monsanto Company, 800 North Lindberg Blvd., D1W, St. 

Louis, MO 63 166 

SCHOTTE, OSCAR E., Department of Biology, Amherst College, Amherst, MA 01002 
SCHUEL, HERBERT, Department of Anatomical Sciences, SUNY, Buffalo, NY 14214 
SCHUETZ, ALLEN W., School of Hygiene and Public Health, Johns Hopkins University, Balti- 
more, MD 2 1205 
SCHWARTZ, JAMES H., Center for Neurobiology and Behavior, New York State Psychiatric 

Institute Research Annex, 722 W. 168th St., 7th Floor, New York, NY 10032 
SCOFIELD, VIRGINIA LEE, Department of Microbiology and Immunology, UCLA School of 

Medicine, Los Angeles, CA 90024 
SEARS, MARY, PO Box 152, Woods Hole, MA 02543 
SEGAL, SHELDON J., Population Division, The Rockefeller Foundation, 1 133 Avenue of the 

Americas, New York, NY 10036 
SELIGER, HOWARD H., Johns Hopkins University, McCollum-Pratt Institute, Baltimore, MD 

21218 
SELMAN, KELLY, Department of Anatomy, College of Medicine, University of Florida, 

Gainesville, FL 32601 

SENFT, JOSEPH, 378 Fairview St., Emmaus, PA 18049 
SHANKLIN, DOUGLAS R., PO Box 1267, Gainesville, FL 32602 
SHAPIRO, HERBERT, 6025 North 13th St., Philadelphia, PA 19141 
SHAVER, GAIUS R., Ecosystems Center, Marine Biological Laboratory, Woods Hole, MA 

02543 

SHAVER, JOHN R., 6 1 5 Jones St., Lansing, MI 489 1 2- 1 7 1 8 
SHEETZ, MICHAEL P., Department of Cell Biology and Physiology, Washington University 

Medical School, 606 S. Euclid Ave., St. Louis, MO 63 1 10 
SHEPARD, DAVID C., PO Box 44, Woods Hole, MA 02543 
SHEPRO, DAVID, Department of Biology, Boston University, 2 Cummington St., Boston, MA 

02215 
SHER, F. ALAN, Immunology and Cell Biology Section, Laboratory of Parasitic Disease, NI- 

AID, Building 5, Room 1 14, NIH, Bethesda, MD 20892 
SHERIDAN, WILLIAM F., Biology Department, University of North Dakota, Grand Forks, ND 

58202 

SHERMAN, I. W., Division of Life Sciences, University of California, Riverside, CA 92502 
SHILO, MOSHE, Department of Microbiological Chemistry, Hebrew University, Jerusalem, 

Israel 

SHOUKIMAS, JONATHAN J., Marine Biological Laboratory, Woods Hole, MA 02543 
SIEGEL, IRWIN M., Department of Ophthalmology, New York University Medical Center, 550 

First Avenue, New York, NY 10016 
SIEGELMAN, HAROLD W., Department of Biology, Brookhaven National Laboratory, Upton, 

NY 11973 
SILVER, ROBERT B., Laboratory of Molecular Biology, University of Wisconsin, 1525 Linden 

Drive, Madison, WI 53706 
SJODIN, RAYMOND A., Department of Biophysics, University of Maryland, Baltimore, MD 

21201 
SKINNER, DOROTHY M., Oak Ridge National Laboratory, Biology Division, Oak Ridge, TN 

37830 
SLOBODA, ROGER D., Department of Biological Sciences, Dartmouth College, Hanover, NH 

03755 



MEMBERS OF THE CORPORATION 25 

SLUDER, GREENFIELD, Cell Biology Group, Worcester Foundation for Experimental Biology, 

22 Maple Ave., Shrewsbury, MA 1 545 

SMITH, MICHAEL A., J 1 Sinabung, Buntu #7, Semarang, Java, Indonesia 
SMITH, PAUL F., PO Box 264, Woods Hole, MA 02543 

SMITH, RALPH I., Department of Zoology, University of California, Berkeley, CA 94720 
SORENSON, MARTHA M., Depto de Bioquimica-RFRJ, Centre de Ciencias da Saude-I.C.B., 

Cidade Universitaria-Fundad, Rio de Janeiro, Brasil 2 1 .9 10 
SPECK, WILLIAM T., Case Western Reserve University, Department of Pediatrics, Cleveland, 

OH 44 106 
SPECTOR, A., College of Physicians and Surgeons, Columbia University, Black Bldg., Room 

1516, New York, NY 10032 

SPEER, JOHN W., Marine Biological Laboratory, Woods Hole, MA 02543 
SPIEGEL, EVELYN, Department of Biological Sciences, Dartmouth College, Hanover, NH 

03755 
SPIEGEL, MELVIN, Department of Biological Sciences, Dartmouth College, Hanover, NH 

03755 
SPRAY, DAVID C., Albert Einstein College of Medicine, Department of Neurosciences, 1300 

Morris Park Avenue, Bronx, NY 10461 

STEELE, JOHN HYSLOP, Woods Hole Oceanographic Institution, Woods Hole, MA 02543 
STEINACHER, ANTOINETTE, Dept. of Otolaryngology, Washington University, School of Med- 
icine, 49 1 1 Barnes Hospital, St. Louis, MO 63 1 10 

STEINBERG, MALCOLM, Department of Biology, Princeton University, Princeton, NJ 08540 
STEPHENS, GROVER C., Department of Developmental and Cell Biology, University of Cali- 
fornia, Irvine, CA 927 1 7 

STEPHENS, RAYMOND E., Marine Biological Laboratory, Woods Hole, MA 02543 
STETTEN, DEWITT, JR., Senior Scientific Advisor, NIH, Bldg. 16, Room 1 18, Bethesda, MD 

20892 

STETTEN, JANE LAZAROW, 2 W Drive, Bethesda, MD 208 14 
STEUDLER, PAUL A., Ecosystems Center, Marine Biological Laboratory, Woods Hole, MA 

02543 

STOKES, DARRELL R., Department of Biology, Emory University, Atlanta, GA 30322 
STOMMEL, ELIJAH W., 766 Palmer Avenue, Falmouth, MA 02540 
STRACHER, ALFRED, Downstate Medical Center, SUNY, 450 Clarkson Ave., Brooklyn, NY 

11203 

STREHLER, BERNARD L., 2235 25 th St., #217, San Pedro, CA 90732 
STRUMWASSER, FELIX, Department of Physiology, Boston University School of Medicine, 

Boston, MA 02 118 

STUART, ANN E., Department of Physiology, Medical Sciences Research Wing 206H, Univer- 
sity of North Carolina, Chapel Hill, NC 275 14 
SUGIMORI, MUTSUYUKI, Department of Physiology and Biophysics, New York University 

Medical Center, 550 First Avenue, New York, NY 10016 
SUMMERS, WILLIAM C., Huxley College, Western Washington University, Bellingham, WA 

98225 
SUSSMAN, MAURICE, Department of Life Sciences, University of Pittsburgh, Pittsburgh, PA 

15260 
SZABO, GEORGE, Harvard School of Dental Medicine, 188 Longwood Avenue, Boston, MA 

02115 
SZENT-GYORGYI, ALBERT, Marine Biological Laboratory, Woods Hole, MA 02543 (deceased 

10/22/86) 
SZENT-GYORGYI, ANDREW, Department of Biology, Brandeis University, Waltham, MA 

02154 
SZENT-GYORGYI, EVA SZENTKIRALY, Department of Biology, Brandeis University, Waltham, 

MA 02 154 
SZUTS, ETE Z., Laboratory of Sensory Physiology, Marine Biological Laboratory, Woods Hole, 

MA 02543 



26 MARINE BIOLOGICAL LABORATORY 

TAMM, SIDNEY L., Boston University Marine Program, Marine Biological Laboratory, Woods 

Hole, MA 02543 

TANZER, MARVIN L., Department of Oral Biology, Medical School, University of Connecti- 
cut, Farmington, CT 06032 
TASAKI, ICHIJI, Laboratory of Neurobiology, Bldg. 36, Rm. 2D10, NIMH, NIH, Bethesda, 

MD 20892 
TAYLOR, DOUGLASS L., Biological Sciences, Mellon Institute, 440 Fifth Avenue, Pittsburgh, 

PA 15213 

TAYLOR, ROBERT E., Laboratory of Biophysics, NINCDS, NIH, Bethesda, MD 20892 
TEAL, JOHN M., Department of Biology, Woods Hole Oceanographic Institution, Woods 

Hole, MA 02543 
TELFER, WILLIAM H., Department of Biology, University of Pennsylvania, Philadelphia, PA 

19174 
THORNDIKE, W. NICHOLAS, Wellington Management Company, 28 State St., Boston, MA 

02109 

TRACER, WILLIAM, Rockefeller University, 1230 York Ave., New York, NY 10021 
TRAVIS, D. M., Veterans Administration Medical Center, Fargo, ND 58102 
TREISTMAN, STEVEN N., Worcester Foundation for Experimental Biology, Shrewsbury, MA 

01545 

TRIGG, D. THOMAS, 1 25 Grove St., Wellesley, MA 02 1 8 1 
TRINKAUS, J. PHILIP, Osborn Zoological Labs, Department of Zoology, Yale University, New 

Haven, CT 065 10 
TROLL, WALTER, Department of Environmental Medicine, College of Medicine, New York 

University, New York, NY 10016 
TROXLER, ROBERT F., Department of Biochemistry, School of Medicine, Boston University, 

80 East Concord St., Boston, MA 02 1 1 8 

TUCKER, EDWARD B., The City University of New York, Baruch College, Box 502, 17 Lexing- 
ton Ave., New York, NY 10010 

TURNER, RUTH D., Mollusk Department, Museum of Comparative Zoology, Harvard Uni- 
versity, Cambridge, MA 02 1 38 
TWEEDELL, KENYON S., Department of Biology, University of Notre Dame, Notre Dame, IN 

46656 
TYTELL, MICHAEL, Department of Anatomy, Bowman Gray School of Medicine, Winston- 

Salem,NC27103 
UENO, HIROSHI, Laboratory of Biochemistry, The Rockefeller University, 1230 York Ave., 

New York, NY 10021 
URETZ, ROBERT B., Division of Biological Sciences, University of Chicago, 950 East 59th St., 

Chicago, IL 60637 
VALIELA, IVAN, Boston University Marine Program, Marine Biological Laboratory, Woods 

Hole, MA 02543 
VALLEE, RICHARD, Cell Biology Group, Worcester Foundation for Experimental Biology, 

Shrewsbury, MA 01 545 

VALOIS, JOHN, Marine Biological Laboratory, Woods Hole, MA 02543 
VAN HOLDE, KENSAL, Department of Biochemistry and Biophysics, Oregon State University, 

Corvallis, OR 97331 
VILLEE, CLAUDE A., Department of Biological Chemistry, Harvard Medical School, Boston, 

MA 02115 
VINCENT, WALTER S., School of Life and Health Sciences, University of Delaware, Newark, 

DE 19711 
WAKSMAN, BYRON, National Multiple Sclerosis Society, 205 East 42nd St., New York, NY 

10017 

WALL, BETTY, 9 George St., Woods Hole, MA 02543 

WALLACE, ROBIN A., Whitney Marine Laboratory, Rte. 1, Box 121, St. Augustine, FL 32086 
WANG, AN, Wang Laboratories, Inc., Bedford Road, Lincoln, MA 01773 
WANG, CHING CHUNG, University of California, School of Pharmacy, San Francisco, CA 

94143 



MEMBERS OF THE CORPORATION 27 

WARNER, ROBERT C, Department of Molecular Biology and Biochemistry, University of Cal- 
ifornia, Irvine, CA 927 1 7 

WARREN, KENNETH S., The Rockefeller Foundation, 1133 Avenue of the Americas, New 
York, NY 10036 

WARREN, LEONARD, Department of Therapeutic Research, School of Medicine, Anatomy- 
Chemistry Building, University of Pennsylvania, Philadelphia, PA 19174 

WATERMAN, T. H., Yale University, Biology Department, Box 6666, 610 Kline Biology 
Tower, New Haven, CT 065 10 

WATSON, STANLEY, Woods Hole Oceanographic Institution, Woods Hole, MA 02543 

WEBB, H. MARGUERITE, Marine Biological Laboratory, Woods Hole, MA 02543 

WEBER, ANNEMARIE, Department of Biochemistry, School of Medicine, University of Penn- 
sylvania, Philadelphia, PA 19174 

WEBSTER, FERRIS, Box 765, Lewes, DE 19958 

WEIDNER, EARL, Department of Zoology and Physiology, Louisiana State University, Baton 
Rouge, LA 70803 

WEISS, LEON P., Department of Animal Biology, School of Veterinary Medicine, University 
of Pennsylvania, Philadelphia, PA 19174 

WEISSMANN, GERALD, New York University, 550 First Avenue, New York, NY 10016 

WERMAN, ROBERT, Neurobiology Unit, The Hebrew University, Jerusalem, Israel 

WESTERFIELD, R. MONTE, The Institute of Neuroscience, University of Oregon, Eugene, OR 
37403 

WEXLER, NANCY SABIN, 1 5 Claremont Avenue, Apt. 92, New York, NY 10027 

WHITE, ROY L., Department of Neuroscience, Albert Einstein College, 1300 Morris Park Ave- 
nue, Bronx, NY 10461 

WHITTAKER, J. RICHARD, Marine Biological Laboratory, Woods Hole, MA 02543 

WIGLEY, ROLAND L., 35 Wilson Road, Woods Hole, MA 02543 

WILSON, DARCY B., Medical Biology Institute, 1 1077 North Torrey Pines Road, La Jolla, CA 
92037 

WILSON, EDWARD O., Museum, Comparative Zoology, Harvard University, Cambridge, MA 
02138 

WILSON, T. HASTINGS, Department of Physiology, Harvard Medical School, Boston, MA 
02115 

WILSON, WALTER L., 743 Cambridge Drive, Rochester, MI 48063 

WITKOVSKY, PAUL, Department of Ophthalmology, New York University Medical Center, 
550 First Ave., New York, NY 10016 

WITTENBERG, JONATHAN B., Department of Physiology and Biochemistry, Albert Einstein 
College, 1 300 Morris Park Ave., New York, NY 1 00 1 6 

WOLFE, RALPH, Department of Microbiology, 1 3 1 Burrill Hall, University of Illinois, Urbana, 
IL61801 

WOODWELL, GEORGE M., 64 Church Street, Woods Hole, MA 02543 (resigned 5/86) 

WORGUL, BASIL V., Department of Ophthalmology, Columbia University, 630 West 168th 
St., New York, NY 10032 

Wu, CHAU HsiUNG, Department of Pharmacology, Northwestern University Medical School, 
203 E. Chicago Ave., Chicago, IL 606 1 1 

WYTTENBACH, CHARLES R., Department of Physiology and Cell Biology, University of Kan- 
sas, Lawrence, KS 66045 

YEH, JAY Z., Department of Pharmacology, Northwestern University Medical School, 303 E. 
Chicago Ave., Chicago, IL 6061 1 

YOUNG, RICHARD W., Mentor O & O, Inc., 3000 Longwater Dr., Norwell, MA 0206 1-1610 

ZACKROFF, ROBERT, 66 White Horn Drive, Kingston, RI 0288 1 

ZIGMAN, SEYMOUR, School of Medicine and Dentistry, University of Rochester, 260 Critten- 
den Blvd., Rochester, NY 14620 

ZIGMOND, RICHARD E., Department of Pharmacology, Harvard Medical School, 250 Long- 
wood Ave., Boston, MA 02 1 1 5 

ZIMMERBERG, JOSHUA J., Bldg. 12A, Room 2007, NIH, Bethesda, MD 20892 

ZOTTOLI, STEVEN J., Department of Biology, Williams College, Williamstown MA 01267 

ZUCKER, ROBERT S., Department of Physiology, University of California, Berkeley, CA 94720 



28 



MARINE BIOLOGICAL LABORATORY 



ASSOCIATE MEMBERS 



ACKROYD, DR. FREDERICK W. 

ADAMS, DR. PAUL 

ADELBERG, DR. AND MRS. EDWARD A. 

AHEARN, MR. AND MRS. DAVID 

ALDEN, MR. JOHN M. 

ALLEN, Miss CAMILLA K. 

ALLEN, DR. NINA S. 

AMON, MR. CARL H. JR. 

ANDERSON, MR. J. GREGORY 

ANDERSON, DRS. JAMES L. AND 

HELENE M. 

ARMSTRONG, DR. AND MRS. SAMUEL C. 
ARNOLD, MRS. Lois 

ATWOOD, DR. AND MRS. KJMBALL C., Ill 
AYERS, MR. DONALD 
BAKER, MRS. C. L. 
BALL, MRS. ERIC G. 
BALLANTINE, DR. AND MRS. H. T., JR. 
BANG, MRS. FREDERIK B. 
BANG, Miss MOLLY 
BANKS, MR. AND MRS. WILLIAM L. 
BARKIN, MR. AND MRS. MEL A. 
BARROWS, MRS. ALBERT W. 
BAUM, MR. RICHARD T. 
BEERS, DR. AND MRS. YARDLEY 
BELESIR, MR. TASOS 
BENNETT, DR. AND MRS. MICHAEL V. L. 
BERG, MR. C. JOHN 
BERNHEIMER, DR. ALAN W. 
BERNSTEIN, MR. AND MRS. NORMAN 
BERWIND, MR. DAVID McM. 
BICKER, MR. ALVIN 
BIGELOW, MRS. ROBERT O. 
BIRD, MR. WILLIAM R. 
BLECK, DR. THOMAS B. 
BOCHE, MR. DAVID 
BODEEN, MR. AND MRS. GEORGE H. 
BOETTIGER, DR. AND MRS. EDWARD G. 

BOETTIGER, MRS. JULIE 

BOLTON, MR. AND MRS. THOMAS C. 

BONN, MR. AND MRS. THEODORE H. 

BORGESE, DR. AND MRS. THOMAS 

BOWLES, DR. AND MRS. FRANCIS P. 
BRADLEY, DR. AND MRS. CHARLES C. 
BRADLEY, MR. RICHARD 
BROWN, MRS. FRANK A., JR. 
BROWN, MR. AND MRS. HENRY 
BROWN, MR. AND MRS. JAMES 
BROWN, MRS. NEIL 
BROWN, DR. AND MRS. THORNTON 
BROYLES, DR. ROBERT H. 
BUCK, DR. AND MRS. JOHN B. 
BUCKLEY, MR. GEORGE D. 
BUNTS, MR. AND MRS. FRANK E. 



BURT, MRS. CHARLES E. 

BUSH, DR. LOUISE 

BUXTON, MR. AND MRS. BRUCE E. 

BUXTON, MR. E. BREWSTER 

CALKINS, MR. AND MRS. G. N., JR. 

CAMPBELL, DR. AND MRS. DAVID G. 

CASE, DR. AND MRS. JAMES 

CARLSON, DR. AND MRS. FRANCIS 

CARLTON, MR. AND MRS. WINSLOW G. 

CHANDLER, MR. ROBERT 

CHASE, MR. TOM H. 

CHILD, DR. AND MRS. FRANK M. 

CHURCH, DR. WESLEY 

CLAFF, MR. AND MRS. MARK 

CLARK, DR. AND MRS. ARNOLD 

CLARK, MR. AND MRS. HAYS 

CLARK, MR. AND MRS. JAMES McC. 

CLARK, MRS. LEONARD B. 

CLARK, MR. AND MRS. LEROY, JR. 

CLARKE, DR. BARBARA J. 

CLEMENT, MRS. ANTHONY 

CLOWES FUND, INC. 

CLOWES, DR. AND MRS. ALEXANDER W. 

CLOWES, MR. ALLEN W. 

CLOWES, DR. AND MRS. G. H. A., JR. 

COBURN, MR. AND MRS. LAWRENCE 
COLEMAN, DR. AND MRS. JOHN 
CONNELL, MR. AND MRS. W. J. 

COOK, DR. AND MRS. PAUL W., JR. 

COPELAND, DR. AND MRS. D. EUGENE 

COPELAND, MR. FREDERICK C. 
COPELAND, MR. AND MRS. PRESTON S. 
COSTELLO, MRS. DONALD P. 
CRABB, MR. AND MRS. DAVID L. 
CRAIN, MR. AND MRS. MELVIN C. 
CRAMER, MR. AND MRS. IAN D. W. 
CRANE, MRS. JOHN O. 
CRANE, JOSEPHINE B., FOUNDATION 
CRANE, MR. THOMAS S. 
CROSS, MR. AND MRS. NORMAN C. 
CROSSLEY, Miss DOROTHY 
CROSSLEY, Miss HELEN 
CROWELL, DR. AND MRS. SEARS 
CURRIER, MR. AND MRS. DAVID L. 
DAIGNAULT, MR. AND MRS. 

ALEXANDER T. 

DANIELS, MR. AND MRS. BRUCE G. 
DAVIDSON, DR. MORTON 
DAVIS, MR. AND MRS. JOEL P. 
DAY, MR. AND MRS. POMEROY 
DECKER, DR. RAYMOND F. 
DEMELLO, MR. JOHN 
DiBERARDiNO, DR. MARIE A. 
DICKSON, DR. WILLIAM A. 



MEMBERS OF THE CORPORATION 



29 



DIEROLF, DR. SHIRLEY H. 

DRUMMEY, MR. AND MRS. CHARLES E. 

DRUMMEY, MR. TODD A. 

DuBois, DR. AND MRS. ARTHUR B. 

DUDLEY, DR. PATRICIA 

DUPONT, MR. A. FELIX, JR. 

DUTTON, MR. RODERICK L. 

EBERT, DR. AND MRS. JAMES D. 

EGLOFF, DR. AND MRS. F. R. L. 

ELLIOTT, MRS. ALFRED M. 

ENOS, MR. EDWARD, JR. 

EPPEL, MR. AND MRS. DUDLEY 

ESTABROOK, MR. GORDON C. 

EVANS, MR. AND MRS. DUDLEY 

FARLEY, Miss JOAN 

FARMER, Miss MARY 

FAULL, MR. J. HORACE, JR. 

FERGUSON, DR. AND MRS. JAMES J., JR. 

FISHER, MRS. B. C. 

FISHER, MR. FREDERICK S., Ill 

FISHER, DR. AND MRS. SAUL H. 

FORBES, MR. JOHN M. 

FORD, MR. JOHN H. 

FRANCIS, MR. AND MRS. LEWIS W., JR. 

FRENKEL, DR. KRYSTINA 

FRIBOURGH, DR. JAMES H. 

FRIENDSHIP FUND 

FRIES, DR. AND MRS. E. F. B. 

FYE, DR. AND MRS. PAUL M. 

GABRIEL, DR. AND MRS. MORDECAI L. 

GAGNON, MR. MICHAEL 

GAISER, DR. AND MRS. DAVID W. 

GALLAGHER, MR. ROBERT O. 

GARFIELD, Miss ELEANOR 

GARREY, DR. WALTER E. 

GELLIS, DR. AND MRS. SYDNEY 

GEPHARD, MR. STEPHEN 

GERMAN, DR. AND MRS. JAMES L., Ill 

GEWECKE, MR. AND MRS. THOMAS H. 

GlFFORD, DR. AND MRS. CAMERON 

GIFFORD, MR. JOHN A. 

GlFFORD, DR. AND MRS. PROSSER 

GILBERT, DRS. DANIEL L. AND CLAIRE 

GILBERT, MRS. CARL J. 

GILDEA, DR. MARGARET C. L. 

GILLETTE, MR. AND MRS. ROBERT S. 

GLAD, MR. ROBERT 

GLASS, DR. AND MRS. H. BENTLEY 

GLAZEBROOK, MR. JAMES 

GLAZEBROOK, MRS. JAMES R. 

GOLDMAN, MRS. MARY 

GOLDRING, MR. MICHAEL 

GOLDSTEIN, DR. AND MRS. MOISE H., JR. 

GOODWIN, MR. AND MRS. CHARLES 

GOULD, Miss EDITH 

GRACE, Miss PRISCILLA B. 



GRANT, DR. AND MRS. PHILIP 
GRASSLE, MRS. J. F. 
GREEN, MRS. DAVIS CRANE 
GREEN, Miss GLADYS M. 
GREER, MR. AND MRS. W. H., JR. 
GRIFFITH, DR. AND MRS. B. HEROLD 
GROSCH, DR. AND MRS. DANIEL S. 
GROSS, MRS. MONA 
GUNNING, MR. AND MRS. ROBERT 
HAAKONSEN, DR. HARRY O. 
HAIGH, MR. AND MRS. RICHARD H. 
HALL, MR. AND MRS. PETER A. 
HALL, MR. WARREN C. 
HALVORSON, DR. AND MRS. HARLYN O. 
HAMSTROM, Miss MARY ELIZABETH 
HARVEY, DR. AND MRS. RICHARD B. 
HASSETT, MR. AND MRS. CHARLES 
HASTINGS, DR. AND MRS. J. WOODLAND 
HAUBRICH, MR. ROBERT R. 
HAY, MR. JOHN 
HAYS, DR. DAVIDS. 
HEDBERG, MRS. FRANCES 
HEDBERG, DR. MARY 
HENLEY, DR. CATHERINE 
HERSEY, MRS. GEORGE L. 
HIATT, DR. AND MRS. HOWARD 
HICHAR, MRS. BARBARA 
HILL, MRS. SAMUEL E. 
HlRSCHFELD, MRS. NATHAN B. 
HOBBIE, DR. AND MRS. JOHN 
HOCKER, MR. AND MRS. LON 
HODGE, MRS. STUART 
HOFFMAN, REV. AND MRS. CHARLES 
HOKIN, MR. RICHARD 
HORNOR, MR. TOWNSEND 
HORWITZ, DR. AND MRS. NORMAN H. 
HOSKIN, DR. AND MRS. FRANCIS C. G. 
HOUSTON, MR. AND MRS. HOWARD E. 
HOWARD, MR. AND MRS. L. L. 
HOYLE, DR. MERRILL C. 

HUETTNER, DR. AND MRS. ROBERT J. 

HUTCHISON, MR. ALAN D. 
HYNES, MR. AND MRS. THOMAS J., JR. 
INOUE, DR. AND MRS. SHINYA 
ISSOKSON, MR. AND MRS. ISRAEL 
JACKSON, Miss ELIZABETH B. 
JAFFE, DR. AND MRS. ERNST R. 
JANNEY, MRS. F. WISTAR 
JEWETT, G. F., FOUNDATION 
JEWETT, MR. AND MRS. G. F., JR. 
JONES, MR. AND MRS. DEWITT C., Ill 
JONES, MR. AND MRS. FREDERICK, II 
JONES, MR. FREDERICK S., Ill 
JORDAN, DR. AND MRS. EDWIN P. 
KAAN, DR. HELEN W. 
KAHLER, MR. AND MRS. GEORGE A. 



30 



MARINE BIOLOGICAL LABORATORY 



KAHLER, MRS. ROBERT W. 
KAMINER, DR. AND MRS. BENJAMIN 
KARPLUS, MRS. ALAN K. 
KARUSH, DR. AND MRS. FRED 
KELLEHER, MR. AND MRS. PAUL R. 
KENDALL, MR. AND MRS. RICHARD E. 
KEOSIAN, MRS. JESSIE 
KEOUGHAN, Miss PATRICIA 
KETCHUM, MRS. PAUL 

KlEN, MR. AND MRS. PlETER 

KINNARD, MRS. L. RICHARD 
KISSAM, MR. WILLIAM M. 
KIVY, DR. AND MRS. PETER 
KOHN, DR. AND MRS. HENRY I. 
ROLLER, DR. LEWIS R. 
KORGEN, DR. BEN J. 
KUFFLER, MRS. STEPHEN W. 
LAFFERTY, Miss NANCY 
LARMON, MR. JAY 
LASTER, DR. AND MRS. LEONARD 
LAUFER, DR. AND MRS. HANS 
LAVIGNE, MRS. RICHARD J. 
LAWRENCE, MR. FREDERICK V. 
LAWRENCE, MR. AND MRS. WILLIAM 
LAZAROW, DR. PAUL 
LEATHERBEE, MRS. JOHN H. 
LEBLOND, MR. AND MRS. ARTHUR 
LEESON, MR. AND MRS. A. Dix 
LEHMAN, Miss ROBIN 
LEMANN, MRS. LUCY B. 
LENHER, DR. AND MRS. SAMUEL 
LEPROHON, MR. JOSEPH 
LEVINE, MR. JOSEPH 
LEVINE, DR. AND MRS. RACHMIEL 
LEVY, MR. STEPHEN R. 
LINDNER, MR. TIMOTHY P. 
LITTLE, MRS. ELBERT 
LIVINGSTONE, MR. AND MRS. ROBERT 
LOEB, MRS. ROBERT F. 

LOVELL, MR. AND MRS. HOLLIS R. 

Low, Miss DORIS 

LOWE, DR. AND MRS. CHARLES W. 

LOWENGARD, MRS. JOSEPH 

MACKEY, MR. AND MRS. WILLIAM K. 
MACLEISH, MRS. MARGARET 
MACNARY, MR. AND MRS. B. GLENN 
MACNlCHOL, DR. AND MRS. 

EDWARD F., JR. 
MAHER, Miss ANNE CAMILLE 
MAHLER, MRS. HENRY 
MAHLER, MRS. SUZANNE 
MANSWORTH, Miss MARIE 
MARSH, DR. AND MRS. JULIAN 
MARTYNA, MR. AND MRS. JOSEPH C. 
MASON, MR. APPLETON 
MASTROIANNI, DR. AND MRS. LUIGI, JR. 



MATHER, MR. AND MRS. FRANK J., III. 
MATHERLY, MR. AND MRS. WALTER 
MATTHIESSEN, DR. AND MRS. G. C. 
McCoY, MRS. Lois 

MCCUSKER, MR. AND MRS. PAUL T. 

MCELROY, MRS. NELLA W. 
MCILWAIN, DR. SUSAN G. 
MCLARDY, DR. TURNER 
MEIGS, MR. AND MRS. ARTHUR 
MEIGS, DR. AND MRS. J. WISTER 
MELILLO, DR. AND MRS. JERRY M. 
MELLON, RICHARD KING, TRUST 
MELLON, MR. AND MRS. RICHARD P. 
MENDELSON, DR. MARTIN 
METZ, DR. AND MRS. CHARLES B. 
MEYERS, MR. AND MRS. RICHARD 
MILLER, DR. DANIEL A. 
MILLER, MR. AND MRS. PAUL 
MIXTER, MR. AND MRS. WILLIAM J., JR. 
MIZELL, DR. AND MRS. MERLE 
MONROY, MRS. ALBERTO 
MONTGOMERY, DR. AND MRS. 

CHARLES H. 
MONTGOMERY, DR. AND MRS. 

RAYMOND B. 
MOOG, DR. FLORENCE 
MOORE, DRS. JOHN AND BETTY 
MORGAN, Miss AMY 
MORSE, MRS. CHARLES L., JR. 
MORSE, DR. M. PATRICIA 
MOUL, DR. AND MRS. EDWIN T. 
MURRAY, DR. DAVID M. 
MYLES-TOCHKO, DR. CHRISTINA J. 
NACE, DR. AND MRS. PAUL 
NACE, MR. PAUL F., JR. 
NELSON, DR. AND MRS. LEONARD 
NELSON, DR. PAMELA 
NEWTON, MR. WILLIAM F. 

NlCKERSON, MR. AND MRS. FRANK L. 

NORMAN, MR. AND MRS. ANDREW E. 
NORMAN FOUNDATION 
NORRIS, MR. AND MRS. BARRY 
NORRIS, MR. AND MRS. JOHN A. 
NORRIS, MR. WILLIAM 
O'HERRON, MR. AND MRS. JONATHAN 
OLSZOWKA, Miss JANICE S. 
O'NEiL, MR. AND MRS. BARRY T. 
C/RAND, MR. AND MRS. MICHAEL 
ORTINS, MR. AND MRS. ARMAND 
O'SULLIVAN, DR. RENEE BENNETT 
PAPPAS, DR. AND MRS. GEORGE D. 
PARK, MRS. FRANKLIN A. 
PARK, MR. AND MRS. MALCOLM S. 
PARMENTER, DR. CHARLES 
PARMENTER, Miss CAROLYN L. 
PELTZ, MR. AND MRS. WILLIAM L. 



MEMBERS OF THE CORPORATION 



31 



PENDERGAST, MRS. CLAUDIA 
PENDLETON, DR. AND MRS. MURRAY E. 
PENNINGTON, Miss ANNE H. 
PERKINS, MR. AND MRS. COURTLAND D. 
PERSON, DR. AND MRS. PHILIP 
PETERSON, MR. AND MRS. E. GUNNAR 
PETERSON, MR. AND MRS. E. JOEL 
PETERSON, MR. RAYMOND W. 
PETTY, MR. RICHARD F. 
PETTY, MR. WILLIAM 
PFEIFFER, MR. AND MRS. JOHN 
PLOUGH, MR. AND MRS. GEORGE H. 
POINTE, MR. ALBERT 
POINTE, MR. CHARLES 
POTHIER, DR. AND MRS. AUBREY 
PORTER, DR. AND MRS. KEITH R. 
PRESS, DRS. FRANK AND BILLIE 
PROSKAUER, MR. RICHARD 
PROSKAUER, MR. JOSEPH H. 
PROSSER, DR. AND MRS. C. LADD 
PSALEDAKIS, MR. NICHOLAS 
PSYCHOYOS, DR. ALEXANDRE 
PUTNAM, MR. ALLAN RAY 
PUTNAM, MR. AND MRS. WILLIAM A., Ill 
RAYMOND, DR. AND MRS. SAMUEL 
REESE, Miss BONNIE 
REINGOLD, MR. STEPHEN C. 
REYNOLDS, DR. AND MRS. GEORGE 
REYNOLDS, MR. ROBERT M. 
REZNIKOFF, MRS. PAUL 
RICCA, DR. AND MRS. RENATO A. 
RIGHTER, MR. HAROLD 
RIINA, MR. AND MRS. JOHN R. 
ROBB, MRS. ALISON A. 
ROBERTS, Miss JEAN 
ROBERTSON, MRS. C. W. 
ROBINSON, DR. DENIS M. 
ROOT, MRS. WALTER S. 

ROSENTHAL, MlSS HlLDE 

ROSLANSKY, DRS. JOHN AND PRISCILLA 

Ross, DR. AND MRS. DONALD 

Ross, DR. ROBERT 

Ross, DR. VIRGINIA 

ROTH, DR. AND MRS. STEPHEN 

ROWE, MR. DON 

ROWE, MR. AND MRS. WILLIAM S. 

RUBIN, DR. JOSEPH 

RUGH, MRS. ROBERTS 

RYDER, MR. AND MRS. FRANCIS C. 

SAGER, DR. RUTH 

SALGUERO, MRS. CAROL G. 

SARDINHA, MR. GEORGE H. 

SAUNDERS, DR. AND MRS. JOHN W. 

SAUNDERS, MRS. LAWRENCE 

SAUNDERS, LAWRENCE, FUND 

SAWYER, MR. AND MRS. JOHN E. 



SAZ, MRS. RUTH L. 

SCHLESINGER, DR. AND MRS. R. WALTER 

SCOTT, DR. AND MRS. GEORGE T. 

SCOTT, MR. AND MRS. NORMAN E. 

SEARS, MR. CLAYTON C. 

SEARS, MR. AND MRS. HAROLD B. 

SEARS, MR. HAROLD H. 

SEAVER, MR. GEORGE 

SEGAL, DR. AND MRS. SHELDON J. 

SENFT, DR. AND MRS. ALFRED 

SHAPIRO, MRS. HARRIET S. 

SHAPLEY, DR. ROBERT 

SHEMIN, DR. AND MRS. DAVID 

SHEPRO, DR. AND MRS. DAVID 

SIMMONS, MR. TIM 

SINGER, MR. AND MRS. DANIEL M. 

SMITH, DRS. FREDERICK E. AND 

MARGUERITE A. 
SMITH, MRS. HOMER P. 
SMITH, MR. VAN DORN C. 
SNYDER, MR. ROBERT M. 
SOLOMON, DR. AND MRS. A. K. 
SPECHT, MRS. HEINZ 
SPIEGEL, DR. AND MRS. MELVIN 
SPOTTE, MR. STEPHEN 
STEELE, MRS. JOHN H. 
STEIN, MR. RONALD 
STEINBACH, MRS. H. BURR 
STETSON, MRS. THOMAS J. 
STETTEN, DR. AND MRS. DEWITT, JR. 
STETTEN, DR. GAIL 
STEWART, MR. AND MRS. PETER 
STONE, MR. ANDREW G. 
STREHLER, DR. AND MRS. BERNARD 
STUNKARD, DR. HORACE 
SUDDITH, MR. WILLIAM 
SWANSON, DR. AND MRS. CARL P. 
SWOPE, MRS. GERARD, JR. 
SWOPE, MR. AND MRS. GERARD L. 

SZENT-GYORGYI, DR. AND MRS. ANDREW 

TABOR, MR. GEORGE H. 

TAYLOR, MR. JAMES K. 

TAYLOR, MRS. MARGERY G. 

TAYLOR, DR. AND MRS. W. RANDOLPH 

TIETJE, MR. AND MRS. EMIL D., JR. 

TIMMINS, MRS. WILLIAM 

TODD, MR. AND MRS. GORDON F. 

TOLKAN, MR. AND MRS. NORMAN N. 

TRACER, MRS. WILLIAM 

TRIGG, MR. AND MRS. D. THOMAS 

TROLL, DR. AND MRS. WALTER 

TUCKER, Miss RUTH 

TULLY, MR. AND MRS. GORDON F. 

ULBRICH, MRS. MARY STEINBACH 

VALOIS, MR. AND MRS. JOHN 

VAN BUREN, MRS. HAROLD 



32 



MARINE BIOLOGICAL LABORATORY 



VAN HOLDE, MRS. KENSAL E. 
VEEDER, MRS. RONALD A. 
VINCENT, DR. WALTER S. 
WAGNER, MR. MARK 
WAKSMAN, DR. AND MRS. BYRON H. 
WARD, DR. ROBERT T. 
WARE, MR. AND MRS. J. LINDSAY 
WARREN, DR. HENRY B. 
WARREN, DR. AND MRS. LEONARD 
WATT, MR. AND MRS. JOHN B. 
WEEKS, MR. AND MRS. JOHN T. 
WEINSTEIN, Miss NANCY B. 
WEISBERG, MR. AND MRS. ALFRED M. 
WEISS, MR. AND MRS. MALCOLM 
WHEELER, DR. AND MRS. PAUL S. 
WHITEHEAD, MR. AND MRS. FRED 
WHITNEY, MR. AND MRS. 
GEOFFREY G., JR. 



WlCHTERMAN, DR. AND MRS. RALPH 
WlCKERSHAM, MR. AND MRS. 
A. A. TlLNEY 

WIESE, DR. CONRAD 

WILHELM, DR. HAZEL S. 

WILSON, MR. AND MRS. T. HASTINGS 

WINN, DR. WILLIAM M. 

WINSTEN, DR. JAY A. 

WITTING, Miss JOYCE 

WOLFINSOHN, MRS. WOLFE 
WOODWELL, DR. AND MRS. GEORGE M. 

YNTEMA, MRS. CHESTER L. 
YOUNG-WALLACE, Miss NINA L. 
ZINN, DR. DONALD J. 
ZIPF, DR. ELIZABETH 
ZWILLING, MRS. EDGAR 



III. CERTIFICATE OF ORGANIZATION 

(On File in the Office of the Secretary of the Commonwealth) 



No. 3 1 70 



We. Alpheus Hyatt, President, William Stanford Stevens, Treasurer, and William T. Sedgwick, 
Edward G. Gardiner, Susan Mims and Charles Sedgwick Minot being a majority of the Trust- 
ees of the Marine Biological Laboratory in compliance with the requirements of the fourth 
section of chapter one hundred and fifteen of the Public Statutes do hereby certify that the 
following is a true copy of the agreement of association to constitute said Corporation, with 
the names of the subscribers thereto: 

We, whose names are hereto subscribed, do, by this agreement, associate ourselves with the 
intention to constitute a Corporation according to the provisions of the one hundred and 
fifteenth chapter of the Public Statutes of the Commonwealth of Massachusetts, and the Acts 
in amendment thereof and in addition thereto. 

The name by which the Corporation shall be known is THE MARINE BIOLOGICAL LABO- 
RATORY. 

The purpose for which the Corporation is constituted is to establish and maintain a laboratory 
or station for scientific study and investigations, and a school for instruction in biology and 
natural history. 

The place within which the Corporation is established or located is the city of Boston within 
said Commonwealth. 

The amount of its capital stock is none. 



In Witness Whereof, we have hereunto set our hands, this twenty seventh day of February in the 
year eighteen hundred and eighty-eight, Alpheus Hyatt, Samuel Mills, William T. Sedgwick, 
Edward G. Gardiner, Charles Sedgwick Minot, William G. Farlow, William Stanford Stevens, 
Anna D. Phillips, Susan Mims, B. H. Van Vleck. 



ARTICLES OF AMENDMENT 33 

That the first meeting of the subscribers to said agreement was held on the thirteenth day of 
March in the year eighteen hundred and eighty-eight. 

In Witness Whereof, we have hereunto signed our names, this thirteenth day of March in the 
year eighteen hundred and eighty-eight, Alpheus Hyatt, President, William Stanford Stevens, 
Treasurer, Edward G. Gardiner, William T. Sedgwick, Susan Mims, Charles Sedgwick Minot. 

(Approved on March 20, 1 888 as follows: 

/ hereby certify that it appears upon an examination of the within written certificate and the 
records of the corporation duly submitted to my inspection, that the requirements of sections 
one, two and three of chapter one hundred and fifteen, and sections eighteen, twenty and 
twenty-one of chapter one hundred and six, of the Public Statutes, have been complied with 
and I hereby approve said certificate this twentieth day of March A.D. eighteen hundred and 
eighty-eight. 

CHARLES ENDICOTT 

Commissioner of Corporations) 

IV. ARTICLES OF AMENDMENT 

(On File in the Office of the Secretary of the Commonwealth) 

We. James D. Ebert, President, and David Shepro, Clerk of the Marine Biological Laboratory, 
located at Woods Hole, Massachusetts 02543, do hereby certify that the following amendment 
to the Articles of Organization of the Corporation was duly adopted at a meeting held on 
August 15, 1975, as adjourned to August 29, 1975, by vote of 444 members, being at least two- 
thirds of its members legally qualified to vote in the meeting of the corporation: 

VOTED: That the Certificate of Organization of this corporation be and it hereby is 
amended by the addition of the following provisions: 

"No Officer, Trustee or Corporate Member of the corporation shall be personally 
liable for the payment or satisfaction of any obligation or liabilities incurred as a 
result of, or otherwise in connection with, any commitments, agreements, activi- 
ties or affairs of the corporation. 

"Except as otherwise specifically provided by the Bylaws of the corporation, meet- 
ings of the Corporate Members of the corporation may be held anywhere in the 
United States. 

"The Trustees of the corporation may make, amend or repeal the Bylaws of the 
corporation in whole or in part, except with respect to any provisions thereof 
which shall by law, this Certificate or the bylaws of the corporation, require action 
by the Corporate Members." 

The foregoing amendment will become effective when these articles of amendment are filed in 
accordance with Chapter 180, Section 7 of the General Laws unless these articles specify, in 
accordance with the vote adopting the amendment, a later effective date not more than thirty 
days after such filing, in which event the amendment will become effective on such later date. 

In Witness whereof and Under the Penalties of Perjury, we have hereto signed our names this 
2nd day of September, in the year 1975, James D. Ebert, President; David Shepro, Clerk. 



34 MARINE BIOLOGICAL LABORATORY 

(Approved on October 24, 1975, as follows: 

I hereby approve the within articles of amendment and, the filing fee in the amount of $10 
having been paid, said articles are deemed to have been filed with me this 24th day of October, 
1975. 

PAUL GUZZI 

Secretary of the Commonwealth) 

V. BYLAWS OF THE CORPORATION OF THE MARINE 
BIOLOGICAL LABORATORY 

(Revised August 16, 1985) 

I. (A) The name of the Corporation shall be The Marine Biological Laboratory. The Cor- 
poration's purpose shall be to establish and maintain a laboratory or sation for scientific study 
and investigation, and a school for instruction in biology and natural history. 

(B) Marine Biological Laboratory admits students without regard to race, color, sex, na- 
tional and ethnic origin to all the rights, privileges, programs and activities generally accorded 
or made available to students in its courses. It does not discriminate on the basis of race, color, 
sex, national and ethnic origin in employment, administration or its educational policies, ad- 
missions policies, scholarship and other programs. 

II. (A) The members of the Corporation ("Members") shall consist of persons elected by 
the Board of Trustees, upon such terms and conditions and in accordance with such proce- 
dures, not inconsistent with law or these Bylaws, as may be determined by said Board of Trust- 
ees. Except as provided below, any Member may vote at any meeting either in person or by 
proxy executed no more than six months prior to the date of such meeting. Members shall 
serve until their death or resignation unless earlier removed with or without cause by the 
affirmative vote of two-thirds of the Trustees then in office. Any member who has attained the 
age of seventy years or has retired from his home institution shall automatically be designated 
a Life Member provided he signifies his wish to retain his membership. Life Members shall 
not have the right to vote and shall not be assessed for dues. 

(B) The Associates of the Marine Biological Laboratory shall be an unincorporated group 
of persons (including associations and corporations) interested in the Laboratory and shall be 
organized and operated under the general supervision and authority of the Trustees. 

III. The officers of the Corporation shall consist of a Chairman of the Board of Trustees, 
President, Director, Treasurer and Clerk, elected or appointed by the Trustees as set forth in 
Article IX. 

IV. The Annual Meeting of the Members shall be held on the Friday following the Second 
Tuesday in August in each year at the Laboratory in Woods Hole, Massachusetts, at 9:30 a.m. 
Subject to the provisions of Article VIII(2), at such meeting the Members shall choose by ballot 
six Trustees to serve four years, and shall transact such other business as may properly come 
before the meeting. Special meetings of the Members may be called by the Chairman or Trust- 
ees to be held at such time and place as may be designated. 

V. Twenty five Members shall constitute a quorum at any meeting. Except as otherwise 
required by law or these Bylaws, the affirmative vote of a majority of the Members voting in 
person or by proxy at a meeting attended by a quorum (present in person or by proxy) shall 
constitute action on behalf of the Members. 



BYLAWS 35 

VI. (A) Inasmuch as the time and place of the Annual Meeting of Members are fixed by 
these Bylaws, no notice of the Annual Meeting need be given. Notice of any special meeting 
of Members, however, shall be given by the Clerk by mailing notice of the time and place and 
purpose of such meeting, at least 1 5 days before such meeting, to each Member at his or her 
address as shown on the records of the Corporation. 

(B) Any meeting of the Members may be adjourned to any other time and place by the 
vote of a majority of those Members present or represented at the meeting, whether or not 
such Members constitute a quorum. It shall not be necessary to notify any Members of any 
adjournment. 

VII. The Annual Meeting of the Trustees shall be held promptly after the Annual Meeting 
of the Corporation at the Laboratory in Woods Hole, Massachusetts. Special meetings of the 
Trustees shall be called by the Chairman, the President, or by any seven Trustees, to be held 
at such time and place as may be designated. Notice of Trustees' meetings may be given orally, 
by telephone, telegraph or in writing; and notice given in time to enable the Trustees to attend, 
or in any case notice sent by mail or telegraph to a Trustee's usual or last known place of 
residence, at least one week before the meeting shall be sufficient. Notice of a meeting need 
not be given to any Trustee if a written waiver of notice, executed by him before or after the 
meeting is filed with the records of the meeting, or if he shall attend the meeting without 
protesting prior thereto or at its commencement the lack of notice to him. 

VIII. (A) There shall be four groups of Trustees: 

( 1 ) Trustees (the "Corporate Trustees") elected by the Members according to such proce- 
dures, not inconsistent with these Bylaws, as the Trustees shall have determined. Except as 
provided below, such Trustees shall be divided into four classes of six, one class to be elected 
each year to serve for a term of four years. Such classes shall be designated by the year of 
expiration of their respective terms. 

(2) Trustees ("Board Trustees") elected by the Trustees then in office according to such 
procedures, not inconsistent with these Bylaws, as the Trustees shall have determined. Except 
as provided below, such Board Trustees shall be divided into four classes of three, one class to 
be elected each year to serve for a term of four years. Such classes shall be designated by the year 
of expiration of their respective terms. It is contemplated that, unless otherwise determined by 
the Trustees for good reason. Board Trustees shall be individuals who have not been considered 
for election as Corporate Trustees. 

(3) Trustees ex officio, who shall be the Chairman, the President, the Director, the Trea- 
surer, and the Clerk. 

(4) Trustees emeriti who shall include any Member who has attained the age of seventy 
years (or the age of sixty five and has retired from his home institution) and who has served a 
full elected term as a regular Trustee, provided he signifies his wish to serve the Laboratory in 
that capacity. Any Trustee who qualifies for emeritus status shall continue to serve as a regular 
Trustee until the next Annual Meeting whereupon his office as regular Trustee shall become 
vacant and be filled by election by the Members or by the Board, as the case may be. The 
Trustees ex officio and emeriti shall have all the rights of the Trustees, except that Trustees 
emeriti shall not have the right to vote. 

(B) The aggregate number of Corporate Trustees and Board Trustees elected in any year 
(excluding Trustees elected to fill vacancies which do not result from expiration of a term) shall 
not exceed nine. The number of Board Trustees so elected shall not exceed three and unless 
otherwise determined by vote of the Trustees, the number of Corporate Trustees so elected 
shall not exceed six. 

(C) The Trustees and Officers shall hold their respective offices until their successors are 
chosen in their stead. 

(D) Any Trustee may be removed from office at any time with or without cause, by vote 
of a majority of the Members entitled to vote in the election of Trustees; or for cause, by vote 



36 MARINE BIOLOGICAL LABORATORY 

of two-thirds of the Trustees then in office. A Trustee may be removed for cause only if notice 
of such action shall have been given to all of the Trustees or Members entitled to vote, as the 
case may be, prior to the meeting at which such action is to be taken and if the Trustee so to 
be removed shall have been given reasonable notice and opportunity to be heard before the 
body proposing to remove him. 

(E) Any vacancy in the number of Corporate Trustees, however arising, may be filled by 
the Trustees then in office unless and until filled by the Members at the next Annual Meeting. 
Any vacancy in the number of Board Trustees may be rilled by the Trustees. 

(F) A Corporate Trustee or a Board Trustee who has served an initial term of at least two 
years duration shall be eligible for re-election to a second term, but shall be ineligible for re- 
election to any subsequent term until two years have elapsed after he last served as Trustee. 

IX. (A) The Trustees shall have the control and management of the affairs of the Corpora- 
tion. They shall elect a Chairman of the Board of Trustees who shall be elected annually and 
shall serve until his successor is selected and qualified and who shall also preside at meetings 
of the Corporation. They shall elect a President of the Corporation who shall also be the Vice 
Chairman of the Board of Trustees and Vice Chairman of meetings of the Corporation, and 
who shall be elected annually and shall serve until his successor is selected and qualified. They 
shall annually elect a Treasurer who shall serve until his successor is selected and qualified. 
They shall elect a Clerk (a resident of Massachusetts) who shall serve for a term of four years. 
Eligibility for re-election shall be in accordance with the content of Article VIII(F) as applied 
to corporate or Board Trustees. They shall elect Board Trustees as described in Article VIII(B). 
They shall appoint a Director of the Laboratory for a term not to exceed five years, provided 
the term shall not exceed one year if the candidate has attained the age of 65 years prior to the 
date of the appointment. They may choose such other officers and agents as they may think 
best. They may fix the compensation and define the duties of all the officers and agents of the 
Corporation and may remove them at any time. They may fill vacancies occurring in any of 
the offices. The Board of Trustees shall have the power to choose an Executive Committee 
from their own number as provided in Article X, and to delegate to such Committee such of 
their own powers as they may deem expedient in addition to those powers conferred by Article 
X. They shall from time to time elect Members to the Corporation upon such terms and condi- 
tions as they shall have determined, not inconsistent with law or these Bylaws. 

(B) The Board of Trustees shall also have the power, by vote of a majority of the Trustees 
then in Office, to elect an Investment Committee and any other committee and, by like vote, 
to delegate thereto some or all of their powers except those which by law, the Articles of Organi- 
zation or these Bylaws they are prohibited from delegating. The members of any such commit- 
tee shall have such tenure and duties as the Trustees shall determine; provided that the Invest- 
ment Committee, which shall oversee the management of the Corporation's endowment funds 
and marketable securities, shall include the Chairman of the Board of Trustees, the Treasurer 
of the Corporation, and the Chairman of the Corporation's Budget Committee, as ex officio 
members, together with such Trustees as may be required for not less than two-thirds of the 
Investment Committee to consist of Trustees. Except as otherwise provided by these Bylaws 
or determined by the Trustees, any such committee may make rules for the conduct of its 
business; but, unless otherwise provided by the Trustees or in such rules, its business shall be 
conducted as nearly as possible in the same manner as is provided by these Bylaws for the 
Trustees. 



X. (A) The Executive Committee is hereby designated to consist of not more than ten 
members, including the ex officio Members (Chairman of the Board of Trustees, President, 
Director and Treasurer); and six additional Trustees, two of whom shall be elected by the 
Board of Trustees each year, to serve for a three-year term. 

(B) The Chairman of the Board of Trustees shall act as Chairman of the Executive Com- 
mittee, and the President as Vice Chairman. A majority of the members of the Executive 



BYLAWS 37 

Committee shall constitute a quorum and the affirmative vote of a majority of those voting at 
any meeting at which a quorum is present shall constitute action on behalf of the Executive 
Committee. The Executive Committee shall meet at such times and places and upon such 
notice and appoint such sub-committees as the Committee shall determine. 

(C) The Executive Committee shall have and may exercise all the powers of the Board 
during the intervals between meetings of the Board of Trustees except those powers specifically 
withheld from time to time by vote of the Board or by law. The Executive Committee may 
also appoint such committees, including persons who are not Trustees, as it may from time to 
time approve to make recommendations with respect to matters to be acted upon by the Execu- 
tive Committee or the Board of Trustees. 

(D) The Executive Committee shall keep appropriate minutes of its meetings and its action 
shall be reported to the Board of Trustees. 

(E) The elected Members of the Executive Committee shall constitute as a standing "Com- 
mittee for the Nomination of Officers," responsible for making nominations, at each Annual 
Meeting of the Corporation, and of the Board of Trustees, for candidates to fill each office as 
the respective terms of office expire (Chairman of the Board, President, Director, Treasurer, 
and Clerk). 

XI. A majority of the Trustees, the Executive Committee, or any other committee elected 
by the Trustees shall constitute a quorum; and a lesser number than a quorum may adjourn 
any meeting from time to time without further notice. At any meeting of the Trustees, the 
Executive Committee, or any other committee elected by the Trustees, the vote of a majority 
of those present, or such different vote as may be specified by law, the Articles of Organization 
or these Bylaws, shall be sufficient to take any action. 

XII. Any action required or permitted to be taken at any meeting of the Trustees, the 
Executive Committee or any other committee elected by the Trustees as referred to under 
Article IX may be taken without a meeting if all of the Trustees or members of such committee, 
as the case may be, consent to the action in writing and such written consents are filed with 
the records of meetings. The Trustees or members of the Executive Committee or any other 
committee appointed by the Trustees may also participate in meeting by means of conference 
telephone, or otherwise take action in such a manner as may from time to time be permitted 
by law. 

XIII. The consent of every 1 rustee shall be necessary to dissolution of the Marine Biologi- 
cal Laboratory. In case of dissolution, the property shall be disposed of in such a manner and 
upon such terms as shall be determined by the affirmative vote of two-thirds of the Board of 
Trustees then in office. 



XIV. These Bylaws may be amended by the affirmative vote of the Members at any meet- 
ing, provided that notice of the substance of the proposed amendment is stated in the notice 
of such meeting. As authorized by the Articles of Organization, the Trustees, by a majority of 
their number then in office, may also make, amend, or repeal these Bylaws, in whole or in part, 
except with respect to (a) the provisions of these Bylaws governing (i) the removal of Trustees 
and (ii) the amendment of these Bylaws and (b) any provisions of these Bylaws which by law, 
the Articles of Organization or these Bylaws, requires action by the Members. 

No later than the time of giving notice of the meeting of Members next following the mak- 
ing, amending or repealing by the Trustees of any Bylaw, notice thereof stating the substance 
of such change shall be given to all Corporation Members entitled to vote on amending the 
Bylaws. 

Any Bylaw adopted by the Trustees may be amended or repealed by the Members entitled 
to vote on amending the Bylaws. 



38 MARINE BIOLOGICAL LABORATORY 

XV. The account of the Treasurer shall be audited annually by a certified public ac- 
countant. 

XVI. Except as otherwise provided below, the Corporation shall, to the extent legally per- 
missible, indemnify each person who is, or shall have been, a Trustee, director or officer of the 
Corporation or who is serving, or shall have served, at the request of the Corporation as a 
Trustee, director or officer of another organization in which the Corporation directly or indi- 
rectly has any interest, as a shareholder, creditor or otherwise, against all liabilities and ex- 
penses (including judgments, fines, penalties and reasonable attorneys' fees and all amounts 
paid, other than to the Corporation or such other organization, in compromise or settlement) 
imposed upon or incurred by any such person in connection with, or arising out of, the defense 
or disposition of any action, suit or other proceeding, whether civil or criminal, in which he or 
she may be a defendant or with which he or she may be threatened or otherwise involved, 
directly or indirectly, by reason of his or her being or having been such a Trustee, director or 
officer. 

The Corporation shall provide no indemnification with respect to any matter as to which 
any such Trustee, director or officer shall be finally adjudicated in such action, suit or proceed- 
ing not to have acted in good faith in the reasonable belief that his or her action was in the best 
interests of the Corporation. The Corporation shall provide no indemnification with respect 
to any matter settled or compromised, pursuant to a consent decree or otherwise, unless such 
settlement or compromise shall have been approved as in the best interests of the Corporation, 
after notice that indemnification is involved, by (i) a disinterested majority of the Board of 
Trustees or of the Executive Committee or, (ii) a majority of the Corporation's Members. 

Indemnification may include payment by the Corporation of expenses in defending a civil 
or criminal action or proceeding in advance of the final disposition of such action or proceeding 
upon receipt of an undertaking by the person indemnified to repay such payment if it is ulti- 
mately determined that such person is not entitled to indemnification under the provisions of 
this Article XVI, or under any applicable law. 

As used in this Article, the terms "Trustee," "director" and "officer" include their respec- 
tive heirs, executors, administrators and legal representatives, and an "interested" Trustee, 
director or officer is one against whom in such capacity the proceeding in question or another 
proceeding on the same or similar grounds is then pending. 

To assure indemnification under this Article of all persons who are determined by the 
Corporation or otherwise to be or to have been "fiduciaries" of any employee benefit plan of 
the Corporation which may exist from time to time, this Article shall be interpreted as follows: 
(i) "another organization" shall be deemed to include such an employee benefit plan, includ- 
ing, without limitation, any plan of the Corporation which is governed by the Act of Congress 
entitled "Employee Retirement Income Security Act of 1974," as amended from time to time 
("ERISA"); (ii) "Trustee" shall be deemed to include any person requested by the Corporation 
to serve as such for an employee benefit plan where the performance by such person of his or 
her duties to the Corporation also imposes duties on, or otherwise involves services by, such 
person to the plan or participants or beneficiaries of the plan; (iii) "fines" shall be deemed to 
include any excise taxes assessed on a person with respect to an employee benefit plan pursuant 
to ERISA; and (iv) actions taken or omitted by a person with respect to an employee benefit 
plan in the performance of such person's duties for a purpose reasonably believed by such 
person to be in the interest of the participants and beneficiaries of the plan shall be deemed to 
be for a purpose which is in the best interests of the Corporation. 

The right of indemnification provided in this Article shall not be exclusive of or affect any 
other rights to which any Trustee, director or officer may be entitled under any agreement, 
statute, vote of members or otherwise. The Corporation's obligation to provide indemnifica- 
tion under this Article shall be offset to the extent of any other source of indemnification or 
any otherwise applicable insurance coverage under a policy maintained by the Corporation or 
any other person. Nothing contained in this Article shall affect any rights to which employees 
and corporate personnel other than Trustees, directors or officers may be entitled by contract, 
by vote of the Board of Trustees or of the Executive Committee or otherwise. 



REPORT OF THE DIRECTOR 39 



VI. REPORT OF THE DIRECTOR 

"The cook was a good cook, as cooks go; and as cooks go she went.' 



Saki 



Transition 



It may be a violation of the best taste, but I believe that I should begin this report, 
my last from the Director's office of the MBL, on a personal note. I do so not because 
personal matters are worthy of the first position, but because a change of the Director- 
ship of the MBL is. Since a good many Corporation Members have only the Annual 
Report in The Biological Bulletin as a source of comprehensive information on the 
Laboratory's work of the prior year, this seems to me the proper place and position 
for a statement on the change. 

I wrote the Director's Report for 1 985-6 in the Candle House. The window before 
which the computer monitor was placed overlooks an angle of Great Harbor and 
Vineyard Sound that I know as well as I know the shape of my hand. This report is 
being written in Charlottesville, Virginia, overlooking Thomas Jefferson's Academi- 
cal Village, the heart of the University of Virginia. In June of 1986 I announced 
to the Executive Committee, and later that summer to the Trustees and the MBL 
community as a whole, that I would be leaving the Directorship and the Laboratory 
at the end of October, 1986. I had accepted, just prior to the first announcement, 
appointment in November as Vice President and Provost at the University of 
Virginia. 

My effort was at the time, as it is now for readers not reached by the earlier one, 
to urge that this decision was made for reasons wholly positive as regards the MBL 
and my relationship with it as of course it has been for Virginia. I found the Labora- 
tory in and no self-congratulation is intended a much stronger position that 
might have been expected at the time of my coming to the Directorship. It now had 
enhanced administrative resources, vastly better physical facilities, a development 
and public relations program that was the envy of larger and richer institutions, 
heightened national and international recognition for the quality of its research and 
instructional programs, and a renewed loyalty of its Corporation membership. 

I had never intended to remain a full-time (or better, double-time) administra- 
torfor that is what, given the limits of my talents, the MBL job quickly became 
beyond the end of a second term, which would have been in 1988. But I had no 
reason to be ashamed of what had been accomplished in nine years of tenure of the 
office. I would have stayed out the term and no more. But the offer from Virginia was 
a very attractive one, and it came from a University I had already served in other 
capacities and for whose intellectual life, Jeffersonian traditions, and physical place I 
have deep admiration. It had always been my intention to return to university life 
after fulfilling a deeply felt obligation to the MBL (where, in reality, I had become a 
biologist). In those circumstances the result of a cost/benefit analysis was clear: no 
great harm would be done to the Laboratory by my leaving it at that time, provided 
that a good transition of leadership could be managed and the Centennial planning 
could be kept on track; and the chances of another university place like as the one at 
Virginia appearing would diminish after 1988. 

Thus it is that I am here and the MBL is there. I miss the Laboratory much more 
than I miss being its Director. There is every reason to believe that, given good health 
and some luck, opportunities to serve it in other ways will arise. I shall take advantage 
of them. 



40 MARINE BIOLOGICAL LABORATORY 

Key to the decision was a judgment as to the effectiveness with which a transition 
to new leadership could be made in 1 986-7. Although I had as yet no way of identify- 
ing the eventual actors therein, my judgment was that the thing could and would be 
done. And so it proved. Fellow embryologist Richard Whittaker agreed generously 
to an infringement of his productive research in order to serve as acting Director. No 
better-qualified person could have been found for this role. 

A distinguished search committee was quickly empaneled, fulfilled its responsibil- 
ity for an exhaustive national search, and was able eventually to recruit as the MBL's 
next Director Harlyn O. Halvorson. His contributions to microbiology are recog- 
nized worldwide, and his activities at the MBL as Trustee, Executive Committee 
Member, Course Director, and advisor to earlier Directors comprise an unmatched 
record of dedicated and effective service. 

The Centennial Committee, already in place at the time of my decision, was obvi- 
ously off to a good start, and its programs for the Centennial year, which begins now 
(August, 1 987), are a fine amalgam of high scientific standards, good taste, and poten- 
tial outreach to a larger public. 

I miss very much, in short, the view out my window from Candle House 301, but 
I feel justified in having had no fears for the health of the institution to which I had 
devoted so much physical and emotional energy in the vicinity of that view. 

Management 

Changing governance and management in an essentially academic institution 
such as the MBL is difficult in principle. It is made more difficult when there is a 
hundred-year history of outstanding achievement; and the difficulties multiply fur- 
ther when the organization is as idiosyncratic (or "horizontal," as Chairman Gifford 
likes to describe it) as is ours. Change was and remains necessary, nevertheless, as I 
have urged in earlier Director's Reports and before what seem to me now numberless 
meetings of Trustees and committees. Nothing that has happened during the last, 
transition-preoccupied year has interfered I am delighted to report with the pro- 
cesses of orderly management change set in motion several years ago. 

The Trustees Board and Corporate have been brought much more closely 
into touch with operations management and decision making than ever before. In- 
deed, this change has accelerated since the last Director's Report was published. The 
new Audit Committee, and other Trustees' committees functioning under revised 
charges, have rendered invaluable service to the Laboratory and its paid administra- 
tion. As a consequence, our financial and political positions have been strengthened 
visibly, even since August of 1986. The Committee on Laboratory Goals, chaired 
conscientiously by Gerald Fischbach and charged a year ago, has produced a short 
but forceful report on those goals on what they should be, and on what steps should 
be taken toward their accomplishment. 

I support their recommendations with enthusiasm. This was an accomplished, 
critical, and independent committee: it is a great pleasure to note that had I written 
the report (to succeed my very solo effort of 1979), it might have been in a different 
prose style, but its content would have been indistinguishable from what is now be- 
fore the MBL community for analysis, debate, and action. 

Treasurer David Currier, who served the Laboratory so handsomely in that posi- 
tion and whose banking skills made possible the splendid new MBL cottage develop- 
ment, retired with the well-earned thanks of the community and has been replaced 
by Robert Manz. Mr. Manz brings to the treasurer's responsibilities more than out- 
standing education, technical, and personal credentials: he is a former officer of Coo- 



REPORT OF THE DIRECTOR 41 

pers and Lybrand, our external auditors for many years, and was a leading member 
of the team assigned to the MBL account. He knows the Laboratory as well as 
perhaps better than any full-time employee. The Treasurer's being another of those 
critical jobs for which the MBL must depend upon volunteers, i.e., upon good will, 
we are doubly lucky to have the good will of Robert Manz. 

Two new management positions were designed, funded, and filled in the course 
of the year. Richard D. Cutler became the Laboratory's first Facilities, Project, and 
Services Manager, and LouAnn D. King accepted appointment as Coordinator of 
Conferences and Housing. These attractive and experienced people have taken on 
challenging responsibilities subdivided and redefined from among the host of such 
carried by former General Manager Homer Smith. 

Mr. Smith's retirement could certainly be described by the cliche, "The End Of 
An Era," and a pretty long era at that. But the description would fail. Homer has left 
his job, to be sure, but he has not left Woods Hole nor the MBL. There is no reason 
to believe that the "era" of his involvement with the Laboratory has ended; and we 
wish him and Cynthia Smith a retirement if that word may be used of continued 
good health and undiminished activity. That, I am sure, is the way they want it to be. 

Donald Ayers, finally, now directs the fully functional Development Office, and 
we can be confident that his programs, aided and additionally monitored by Lisa 
Thimas (formerly Assistant to the Director), will flourish during the Centennial Year 
before us. 

Remarkably, these are a mere sample, not the totality of appointments and 
change processes that took place during the transition year. Let no one be concerned 
about a let-down of effort or a diminution of those management skills of which the 
MBL has been so much in need. All that we need to be concerned about is the means 
of increasing further the rate of positive change. 

Systems 

In that connection it is noteworthy that three of the critical systems upon which 
those who must operate, and make decisions for, the MBL have been changed and 
improved during the year past. Most important, the many roadblocks, internal as 
well as external, impeding progress toward a rational system of overhead recovery 
have been eliminated. The system is now in operation, and although it works no 
better than those of our peer institutions, it is no worse. The high optimism implicit 
in such a negative statement will be understood by all who have grappled in decades 
past with the problem of reimbursing the Laboratory for its costs in housing and 
supporting research. 

A job classification system was designed, discussed widely, and put in place for all 
MBL employees. This had been the majority wish for many years, and its fulfillment 
has indeed brought a measure of regularity and central accountability to the manage- 
ment of operations and personnel. The system includes not only a set of objective 
job descriptions and grades, but also processes for the hearing and adjudication of 
disputes and grievances. Already called upon for service in that connection has been 
the Classification Review Board established for the purpose and chaired, ably as al- 
ways, by Joan Howard. 

Last but not least, the Controller's department now has a greatly improved system 
of accounts and data management, aided by the appointment of an Assistant Control- 
ler with excellent accounting skills and a resident specialist in electronic data process- 
ing. It is not amiss to note that these changes are a direct consequence of the enhanced 
Trustee oversight initiated two years ago, to which I have referred above. 



42 MARINE BIOLOGICAL LABORATORY 

Research and education 

Summer research activities and accomplishments were at their now accustomed 
high levels in the summer of 1986 and, as will be evident to readers of this report by 
the time it has been published, so will they be in 1 987. There are very few laboratories 
or library accommodations unspoken for in summer at the MBL. The only concern 
I have heard expressed from time to time is that not enough applicants are rejected, 
or, to put it in the way it is usually phrased, that the MBL ought to have more of a 
choice among potential summer scientists. Perhaps so; but the financial realities and 
demography of those disciplines practiced by MBL summer investigators speak 
differently to me. I believe that we do very well to have a significant percentage of the 
most honored neuroscientists, cell biologists, developmental biologists, microsco- 
pists, and the like here at the MBL every summer, and it is insurance against stagna- 
tion that we can admit most of those others, not yet so honored perhaps, who have 
legitimate research to perform here. And lest the reader believe what is sometimes 
unfairly and untruthfully asserted, that the MBL is a club for its "regulars" and 
nothing more, let him inspect the roster of last summer's MBL Fellows and indepen- 
dent investigators for the new and the still-young. The evidence is heartening. No 
other organization can boast so large and so diverse an assembly of biologists engaged 
upon serious research for a meaningful part of the year. 

The year-round research program, focus of the plan presented by the Committee 
on Laboratory Goals, ended the year in about the same configuration as at the begin- 
ning. It was as large an enterprise as the MBL can accommodate within existing build- 
ings; it was well-funded; and it was steadily productive. As I have said elsewhere, there 
is no university for which such a group, collected together as a Biology Department, 
would not be a prize. 

There are, however, changes imminent. The National Institute for Neurological 
and Communicative Diseases and Stroke has decided, for what are considered in 
Bethesda to be good reasons, on an eventual recall of at least the bulk of its on- 
location program at the MBL. It is possible that a part of this effort will remain for 
several years, but under the existing directives, most of the program will revert to 
permanent residence on the NIH campus, with summer research at the MBL. Balanc- 
ing this, there have been selections from among excellent applicant investigators for 
year-round accommodation, so that the size of the resident research program will not 
change very much in the near term, nor will its very high quality overall. 

If, however, the Corporation and Trustees elect to implement the recommenda- 
tions of the Fischbach committee, there will be major change indeed, and it will have 
to be accompanied by new methods of funding and supporting year-round scientists, 
new programs, a net increase in size, and large additions to the inventory of research 
space and general-use facilities. 

The instructional program, whose unique contribution to the scientific manpower 
of this nation has been recognized by astonishingly (for these times) generous support 
from government and private agencies, fared well in 1 986 and will clearly continue 
thus in 1987. In no small part this is due to the far-sighted support of such private 
donors as the Markey Trust, the Pew Memorial Trust, the Grass Foundation, and 
the Klingenstein Fund. But success and a tough-minded maintenance of the highest 
standards of quality are also testimony to the very spirit of the MBL, inherited and 
still vigorous, as established in its teaching programs by the Founders. In no small 
measure the smooth operation of our complex instructional enterprise much more 
complex, in important ways, than the mounting of courses in a university has de- 
pended, and will probably continue to depend, upon three things: ( 1 ) the flow of 
financial support for indirect as well as direct costs from private donors; (2) the collab- 



REPORT OF THE DIRECTOR 43 

oration, especially via equipment loans, of the world's leading manufacturers of sci- 
entific instruments; and (3) wise, artful, and minimally intrusive management of the 
courses and their people, as exemplified in the work of Joan Howard and her ever- 
helpful staff in the Office of Sponsored Programs. 

Successful a story as this is, it will not remain so without continuous effort. It was 
a principle of the last administration, as it was of the MBL's first, that the instructional 
program shares the first priority with research, and that the teaching is as much on 
the moving frontier of our science as is the research of any particular laboratory. The 
principle will continue, I hope, to be held. Thus far the signs are good. By way of 
example, I might cite an unusual and already noteworthy collaboration of the MBL 
and the University of Georgia in the teaching of Plant Cell and Molecular Biology. 
And if the current plans for the next version of Embryology are implemented, it will 
be a further signal to the effect that curricular and organizational adventurousness 
need not be absent from teaching by an eminent faculty, to the world's most able 
classes of young biologists. 

If all the above has the scent of hyperbole, I apologize but also deny any such 
purpose. The MBL is a very remarkable organism: the bare facts upon which this 
summary is based are printed in the accompanying pages of the Annual Report and 
in other MBL publications. I have simply written a summary, with a few laudatory 
adjectives where they are clearly justified. We tend easily to forget, as intimates of 
quality, how special a thing it is in the world. We do not forget problems with the 
same celerity: those, like the crying of a baby, evolve a sound that cannot be ignored. 

Albert Szent-Gyorgyi 

Among the score of Nobelists associated with the Laboratory during the two or 
three decades past, none made such a mark upon the day-to-day life of the place as 
did Albert Szent-Gyorgyi, who died at a ripe old age during the year. Of course that 
mark was to some extent a consequence of Szent-Gyorgyi's year-round residence in 
Woods Hole and at the MBL, from a time when his Institute for Muscle Research 
was virtually the only active group during the non-summer months until a few years 
ago, when it was well-eclipsed, in size and in visibility, by other year-round labora- 
tories. But his influence did not depend solely upon presence. Prof's was a mind and 
a personality of most unusual strength. His intellectual power and personal charm 
were merely facets of the whole man, who was also possessed of moral power, a re- 
markable persuasiveness, an abundance of love and good cheer, and overlying all, an 
aesthetic imperative that drove him to the still-unexplored for its beauty first, and is 
utility second. During the height of his career, when I was a student here, he was for 
many of us the paradigm of the scientist as the spearhead of culture. Throughout his 
long life, even at the very end, when he and I could converse only in shouts, he was 
for me and, I know, for many others, the very model of a man. 

Coda 

The MBL nurtures us all, as men and women, as extenders of culture, as investiga- 
tors, as teachers of the best science to the best students. Long may it flourish; and to 
it a happy hundredth birthday! 

VII. REPORT OF THE TREASURER 

This is my first report to you as Treasurer. Although I have been at the job for 
almost a year now, this is my first chance to say that I am privileged and honored to 
have an opportunity to contribute to this excellent and exciting institution. I look 



44 MARINE BIOLOGICAL LABORATORY 

forward to working with the staff, Executive Committee, Trustees, and Corporation 
as we move into the Laboratory's Second Century. 

The financial statements of the Laboratory for the year ended December 31,1 986, 
follow this report. Before I comment on the financial results for the year, I must 
discuss some changes in presentation that have been made to the financial statements 
to reflect actions taken by the Corporation and the Executive Committee in the last 
year and to more clearly present the financial position of the Laboratory. 

In the Statement of Support, Revenues, Expenses, and Changes in Fund Balances 
(Statement of Support), a separate column has been provided for the newly estab- 
lished Housing Enterprises Fund within the Current Unrestricted Fund. The Balance 
of Operations for this fund of $ 1 12,294 has been transferred to a Reserve for Repairs 
and Replacements restricted to expenditure on housing. This is a major step forward 
for the Laboratory in providing for its financial future; I applaud the wisdom of your 
decision to establish this fund. 

The financial statements no longer show the assets or results of operations of the 
Retirement Funds. These funds are not actually assets of the MBL but are owned by 
the Laboratory's pension plan, so they should not appear in the Laboratory's financial 
statements but in those of the Plan. To date there have been no separately issued 
financial statements of the Retirement Plan because of its size. The Plan has now 
grown large enough that provisions of the Employees' Retirement Income Security 
Act require separate financial statements of the Plan; these have been prepared, and 
the Retirement Fund has been removed from the Laboratory's financial statements. 

Also affecting pension accounting, the Laboratory has adopted the new require- 
ments of the Financial Accounting Standards Board on accounting for pension ex- 
pense of the Laboratory (i.e., its obligation to contribute to the plan). The new re- 
quirements seek to more accurately reflect the impact of changes in investment and 
annuity market conditions as well as employment and actuarial expectations of the 
employer's pension obligation. For the Laboratory this has meant a reduction in pen- 
sion expense from $ 1 42,833 in 1985 to $82,682 in 1 986, chiefly because of the favor- 
able performance of the Retirement Plan's investment portfolio. 

The Balance Sheet now reflects the market value of the Laboratory's invested 
funds, rather than their book value. I have recommended this change so that you will 
always have before you the actual market value of the endowment rather than a book 
number which reflects only the timing of past donations and the results of past invest- 
ment activity. 

I trust that these comments will assist you as you examine the Laboratory's Finan- 
cial statements. 

The results of operations for 1986 show a picture of continued current operating 
strength, with some fluctuations that bear watching. They suggest some significant 
improvements in the long-term strength of the Laboratory if trends begun this year 
can be sustained and expanded and they point to an agenda for future actions. 

Total support and revenues increased from 1985 to 1986 by $272,488 while ex- 
penses increased by $269,648. 

Within support and revenues, gains in gifts, recovery in indirect costs from the 
summer program (laboratory fees), and dormitory and dining fees were offset by de- 
clines in direct support for year-round research and the associated recovery of indirect 
costs, and slight declines in the income of Research Services and Marine Resources. 
It is encouraging to see some strength in fee income it suggests that the Laboratory 
renders a measurable service which can be directly supported by the users of that 
service but for the same reason it is discouraging to see the declines in Research 
Services and Marine Resources income. I have recommended to the Director that a 



REPORT OF THE TREASURER 45 

careful examination of all the fee for service income of the Laboratory be made in 
order to make the best possible match between service and income, and to guarantee 
the future economic viability of our services. 

Gifts increased by more than $200,000 in 1986. We have continued to receive 
strong support from the Pew and Markey foundations for the instructional program; 
we made significant strides in the Mellon match gift for Library endowment; and 
we received a $200,000 gift from the Monsanto Corporation in anticipation of our 
Centennial celebration. Such dedicated support allows your laboratory to maintain 
the excellence of its programs. As your Treasurer, I must point out, however, that we 
are perilously dependent on the generosity of donors of gifts for current use. In order 
to achieve the assurance of continued excellent programs, we need to seek a dramatic 
increase in our endowment. 

Within the expense categories, I call your attention to the increases in housing and 
depreciation expense, that in aggregate, amount to almost $250,000. Of this amount, 
approximately $133,000 is attributable to the additional housing units at Memorial 
Circle and an additional $76,000 was expensed as administrative costs, thus complet- 
ing our planned "full costing" of the separate Housing Enterprises Fund. More sig- 
nificantly, if you look down the column entitled "Housing Enterprises Fund," you 
will note that for the first time, the MBL, as projected and previously mentioned, has 
funded $ 1 12,294 in depreciation costs associated with the housing enterprise. These 
funds are being set aside to help finance major future capital improvements in the 
housing facilities. 

We again ended the year with an excess of revenues over expenses in the current 
unrestricted fund ($70,590). You should thank your Controller, John Speer, for his 
role in achieving this result. The Laboratory is well served by John's vigilance over 
the operating budget and his ability to see clouds on the financial horizon and recom- 
mend a course long before the storm strikes. 

As your Treasurer, I will never use the word "surplus" in connection with an 
excess of revenues over expenses until the Laboratory has been able to use that excess 
to fund depreciation on its plant. As I told the Trustees last winter and hope to dem- 
onstrate to the rest of the Corporation, the annual "surpluses" of the Laboratory are 
wiped out when depreciation is taken into account. We have in fact been able to 
maintain the quality of our physical plant through the generosity of our donors, but 
this means that the heroic capital fund raising efforts of our directors in the last ten 
years have been required to maintain rather than to improve the quality of our plant. 

I believe we must set as a goal the funding of a significant portion of our deprecia- 
tion expense from operating revenues. This year we took a modest first step towards 
that goal and transferred $33,650 from the current unrestricted fund to a reserve for 
replacements. We must do much more in the future. 

In this report I have tried to indicate those aspects of our financial condition that 
merit our attention and will require concerted action. I have no doubt left some of 
your questions on the financial performance of 1986 unanswered here, but I welcome 
them directly and will do my best to respond. 



46 MARINE BIOLOGICAL LABORATORY 



certified public accountants One Post Office Square in principal areas ot the world 

Boston, Mass 02109 

telephone (617) 574-5000 
TWX 710-321-0489 
telex 6817018 



&Lybrand 



To the Trustees of 

Marine Biological Laboratory 

Woods Hole, Massachusetts 

We have examined the balance sheet of Marine Biological 
Laboratory as of December 31, 1986 and the related statement of support, 
revenues, expenses and changes in fund balances for the year then ended. 
Our examination was made in accordance with generally accepted auditing 
standards and, accordingly, included such tests of the accounting records 
and such other auditing procedures as we considered necessary in the 
circumstances. We previously examined and reported upon the financial 
statements of the Laboratory for the year ended December 31 1985, which 
condensed statements are presented for comparative purposes only. 

In our opinion, the financial statements referred to above 
present fairly the financial position of Marine Biological Laboratory at 
December 31, 1986 and its support, revenues, expenses and changes in fund 
balances for the year then ended, in conformity with generally accepted 
accounting principles applied on a basis consistent with that of the 
preceding year, except for the changes, with which we concur, in the 
method of accounting for investments as described in Note C, the method of 
accounting for pension expense as described in Note E and the method of 
accounting for pension funds as described in Note J. 

Very truly yours, 

o 

Boston, Massachusetts VOOOCTO f, 

April 18, 1987 




REPORT OF THE TREASURER 



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50 MARINE BIOLOGICAL LABORATORY 

NOTES TO FINANCIAL STATEMENTS 

A. Purpose of the Laboratory: 

The purpose of Marine Biological Laboratory (the "Laboratory") is to establish and maintain a 
laboratory or station for scientific study and investigations, and a school for instruction in biology 
and nature history. 

B. Significant Account ing Policies: 

Basis of Presentation Fund A ccounting 

In order to ensure observance of limitations and restrictions placed on the use of resources available 
to the Laboratory, the accounts of the Laboratory are maintained in accordance with the principles 
of "fund accounting." This is the procedure by which resources are classified into separate funds in 
accordance with specified activities or objectives. 

Externally restricted funds may only be utilized in accordance with the purposes established by the 
donor or grantor of such funds. However, the Laboratory retains full control over the utilization of 
unrestricted funds. Restricted gifts, grants, and other restricted resources are accounted for in the 
appropriate restricted funds. Restricted current funds are reported as revenue when received and as 
related costs are incurred. Unrestricted current funds are reported as revenue when earned. 

Endowment funds are subject to restrictions requiring that the principal be invested with income 
available for use for restricted or unrestricted purposes by the Laboratory. Quasi-endowment funds 
have been established by the Laboratory for the same purposes as endowment funds; however, the 
principal of these funds may be expended for various restricted and unrestricted purposes. 

Fixed Assets 

Fixed assets are recorded at cost. Depreciation is computed using the straight-line method over esti- 
mated useful lives of fixed assets. 

Reclassifications 

The financial statements for 1986 reflect certain changes in classification of revenue, expenses and 
changes in fund balances. Similar reclassifications have been made to amounts previously reported 
in order to provide consistency of the financial statements. In addition, the financial statements 
reflect in 1986 the segregation of the current unrestricted fund balance into two components: the 
current unrestricted and the housing enterprise fund balances. 

Contracts and Grants 

Revenues associated with contracts and grants are recognized in the statement of support, revenues, 
expenses and changes in fund balances when received and as related costs are incurred. The Labora- 
tory reimbursement of indirect costs relating to government contracts and grants is based on negoti- 
ated indirect cost rates with adjustments for actual indirect costs in future years. Any over- or under- 
recovery of indirect costs is recognized through future adjustments of indirect cost rates. 



Investments 

Investments purchased by the Laboratory are carried at market value (Note C). Money market secu- 
rities are carried at cost which approximates market value. Investments donated to the Laboratory 
are carried at fair market value at the date of the gift. For determination of gain or loss upon disposal 
of investments, cost is determined based on the average cost method. The Laboratory is the benefi- 
ciary of certain endowment investments which are held in trust by others. These investments are 
reflected in the financial statements. Every ten years the Laboratory's status as beneficiary is reviewed 
to determine that the Laboratory's use of these funds is in accordance with the intent of the funds. 



REPORT OF THE TREASURER 5 1 

Investment Income and Distribution 

The Laboratory follows the accrual basis of accounting except that investment income is recorded 
on a cash basis. The difference between such basis and the accrual basis does not have a material 
effect on the determination of investment income earned on a year-to-year basis. 

Investment income includes income from the investments of specific funds and from the pooled 
investment account. Income from the pooled investment account is distributed to the participating 
funds on the basis of their proportionate share at market value adjusted for any additions or disposals 
to pooled funds. 

C. Change in Accounting Met hod for Investments: 

Effective January 1, 1986, the Laboratory adopted the accounting policy of reporting investments 
and the related fund balances at market value to more clearly reflect the financial impact of the 
Laboratory's investment policies. Investments and the related fund balances in prior years were 
reported at cost. The cumulative increase in the fund balances at December 31,1 986 and 1 985 is as 
follows: 

Current Restricted Funds: 1986 1 985 

Unexpended gifts $ 11,782 

Endowment funds: 

Unrestricted $ 80,070 329,166 

Restricted 81,780 215,763 

Quasi-endowment funds: 

Unrestricted 31,438 126,333 

Restricted 142,527 436,913 

Increase in unrealized appreciation and related 

fund balances $335,815 $1.119,957 

This change has been retroactively applied to the fund balances as of the beginning of the year ended 
December 3 1 , 1985 as follows: 

Unexpended gifts $ (364) 

Endowment funds: 

Unrestricted 263,969 

Restricted 311,892 



Quasi-endowment funds: 

Unrestricted (3,747) 

Restricted 104.458 

Cumulative unrealized gain/loss $ 676,208 



D. Land, Buildings, and Equipment: 

The following is a summary of the unrestricted plant fund assets: 

1986 1985 

Land $ 689,660 $ 689,660 

Construction in progress 140,826 

Buildings 16,333,358 14,861,244 

Equipment 2,170,878 2.113.321 

19,193,896 17,805,051 

Less accumulated depreciation (7,143.565) (6,579.654) 

$12,050,331 $11,225,397 



52 



MARINE BIOLOGICAL LABORATORY 



E. Retirement Fund: 

During 1986, the Laboratory elected early application of Statement of Financial Accounting Stan- 
dard No. 87, "Employer's Accounting for Pensions." This Statement establishes standards of finan- 
cial accounting and reporting for an employer that offers pension benefits to its employees and super- 
cedes earlier standards. The early election reduced the actuarially determined pension expense from 
$132,866 to $82,682. 

The Laboratory has a noncontributory defined benefit pension plan for substantially all employees. 
Contributions are intended to provide for benefits attributed to the service date, but also those ex- 
pected to be earned in the future. 



Actuarial present value of benefit obligations: 
Accumulated benefit obligation including vested benefits of 
$1,484,283 

Projected benefit obligation 

Plan assets at fair value 

Projected benefit obligation less than plan assets 

Unrecognized net (gain) or loss 

Prior service cost not yet recognized in net periodic pension cost 

Unrecognized net obligation at March 1, 1986 

Prepaid pension cost (pension liability) recognized in the statement 
of financial position 

Net pension cost for fiscal year ending December 31,1 986: 
Service cost benefits earned during the period 
Interest cost on projected benefit obligation 
Actual return on plan assets 
Net amortization and deferral 

Net periodic pension cost 



1,686.685 

2.561,619 

2.608,987 

47,368 

186,054 

(316.104) 
$ (82.682) 

138,391 

142,381 

(319,877) 

121.787 

$ 82,682 



The actuarial present value of the projected benefit obligation was determined using a discount rate 
of 7.3% and rates of increase in compensation levels of 6%. The expected long-term rate of return 
on assets was 8%. 

In addition, the Laboratory participates in the defined contribution pension program of the Teachers 
Insurance and Annuity Association. Expenses amounted to $106,535 in 1986 and $95,858 in 1985. 

F. Pledges and Grants: 

As of December 31, 1986 and 1985, the following amounts remain to be received on gifts and grants 
for specific research and instruction programs, and are expected to be received as follows: 



December 31. 1986 



December 31, 1985 



1987 
1988 
1989 



Unrestricted 

$20,000 
10,000 



$30,000 



Restricted 

$615,027 

40,764 

5.764 

$661,555 



Unrestricted 

$10,000 
10,000 



$20,000 



Restricted 

$550,240 
15,000 



$565.240 



G. Interfund Borrowings: 

Interfund balances at December 3 1 are as follows: 

Current Funds 

Due to plant funds 

Due to endowment funds 

Due to restricted quasi-endowment funds 



1986 



$(169,615) 
(115,909) 
(200.750) 

$(486,274) 



1985 

$ (56,669) 
(156,622) 

$(213,291) 



REPORT OF THE TREASURER 53 

H. Mortgages and Notes Payable: 

The mortgage note payable with a term of 26 years is in the amount of $ 1 .3 million bearing interest 
based on the bank's prime rate plus three quarters percent (.75%) on a floating basis for the initial 
five year period with a floor of 7.50% and a ceiling of 1 3.00%. The interest rate at December 31,1 986 
was 9.00%. The mortgage loan is collateralized by a first mortgage on the land and properties known 
as Memorial Circle, with recourse in the event of default limited to this land and property and the 
related revenue. Principal and interest payments of $ 1 5,000 are due and payable monthly commenc- 
ing January 19, 1987. 

Other notes payable consist of the following: 

Unsecured note with interest at 7.90% with monthly principal 
payments of $22 1.20 plus interest $ 9,194 

Unsecured note with interest at 6.90% with monthly principal 
payments of $394. 71 plus interest 12.990 

$22,184 



At December 31, 1986, these mortgages and notes payable had aggregate future annual principal 
payments as follows: 

Amount 

1987 $ 73,001 

1988 78.654 

1989 83.899 

1990 86,492 

1991 94,127 
Thereafter 906.010 

1,322,183 

Less current portion 73.001 

$1,249,182 



54 



MARINE BIOLOGICAL LABORATORY 



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REPORT OF THE LIBRARIAN 55 

VIII. REPORT OF THE LIBRARIAN 

Our serial titles are now all entered into the On-Line OCLC catalog which is based 
in Ohio. The number of requests for copies of articles in our periodical collection has 
doubled since this project was completed. Over 350 libraries and laboratories sent 
requests during 1986, and we now serve a larger scientific community than we did 
before our collection was included in this database. 

In preparation for the Centennial we have prepared a computer program for the 
records in the Archives. Lists of all scientists, students, lecturers, employees, and oth- 
ers who have been at the MBL since 1 888 will be placed in an archival database. This 
information will be valuable to science historians, our public information office, and 
the library reference staff. Most of this material will be entered by Ruth Davis and 
her volunteer staff. Photographs held in the Rare Books and Archives area are being 
cataloged. Negatives are being made of a number of the rare ones. Many of these 
photographs will be used in a Centennial book that is being planned for 1988. 

We gave one-day tours of the Library in June to two groups of librarians. One was 
a group from the Boston meeting of the Special Librarians Association and the other 
was a group named the "Rte. 1 28 Librarians" from the Hi-Tech libraries in that area. 

Binding increased this year since we picked up a number of volumes where one 
or two issues were missing. We bound these noting the "lacks" on the spine. Over 
3000 volumes were sent to the binders during the winter months. 

IX. EDUCATIONAL PROGRAMS 



SUMMER 

BIOLOGY OF PARASITISM 

Course directors 

ENGLUND, PAUL, Johns Hopkins University 
SHER, ALAN, NIAID/NIH 

Other faculty, staff, and lecturers 

ALDRITT, SUSAN, Harvard University 

BEVERLY, STEPHEN, Harvard Medical School 

BLOOM, BARRY, Albert Einstein College of Medicine 

BROWN, KIM, University of Iowa 

BURAKOFF, STEVEN, J., Dana-Farber Cancer Institute 

BURNS, JAMES, Hahnemann University 

BUTTERWORTH, ANTHONY, University of Cambridge, UK 

CANTOR, CHARLES, Columbia University 

CARTER, RICHARD, NIAID/NIH 

CERAMI, ANTHONY, Rockefeller University 

CLEVELAND, DON, Johns Hopkins University 

DINTZIS, HOWARD, Johns Hopkins University 

DONELSON, JOHN, University of Iowa 

DOOLITTLE, RUSSELL, University of California, San Diego 

DVORAK, JAMES, NIAID/NIH 

GEARHART, PATRICIA, Johns Hopkins University 

GOTTLIEB, MICHAEL, Johns Hopkins University 

HART, GERALD W., Johns Hopkins University 

HERELD, DALE, Johns Hopkins University 

HOWARD, JAMES, Wellcome Laboratories 



56 MARINE BIOLOGICAL LABORATORY 

HOWARD, RUSSELL, NIAID/NIH 

JAMES, STEPHANIE, George Washington School of Medicine 

JOINER, KEITH, NIAID/NIH 

KNOPF, PAUL, Brown University 

MARTINEZ-PALOMO A., Center for Advanced Research 

McMAHON-PRATT, D., Yale University Medical School 

Moss, BERNARD. NIAID/NIH 

NELSON, GEORGE, University of Liverpool, UK 

NEVA, FRANKLIN A., NIAID/NIH 

NUSSENZWEIG, VICTOR, New York University Medical Center 

OTTESON, ERIC, NIAID/NIH 

PEREIRA, MIERCIO, Tufts University School of Medicine 

PFEFFERKORN, ELMER, Dartmouth Medical School 

VAN DER PLOEG, LEX, Columbia University 

RIBEIRO, JOSE, Harvard University 

ROCK, THEODORE, Howard Hughes Medical Institute 

SACKS, DAVID, NIAID/NIH 

SCOTT, PHILLIP, NIAID/NIH 

SHARKEY, ANDREW, University of Edinburgh, UK 

SHEVACH, ETHAN, NIAID/NIH 

SPIELMAN, ANDREW, Harvard University School of Public Health 

STRAND, METTE, Johns Hopkins University 

SUPLICK, KATHY, Hahnemann University 

TURNER, MERVYN J., Merck Sharp and Dohme Research Laboratories 

WALLIKER, D., University of Edinburgh, UK 

WANG, CHING C., University of California, San Francisco 

WARD, DAVID, Yale University 

WARD, SAMUEL, Carnegie Institute 

WARREN, KENNETH, Rockefeller Foundation 

WASSOM, DONALD, University of Wisconsin 

Students 

ALANO, PIETRO, University of Milan, Italy 

ANDERSEN, BIRGITTE JYDING, Statens Serum Institute, Denmark 

DOSHI, PARULD., UMDNJ-Rutgers Medical School 

EID, JOSIANE E., Johns Hopkins University 

HARYANA, SOFIA M., Gadjah Mada University, Indonesia 

LOMBARDI, GEORGE V., Washington University 

Lucius, RICHARD H. C., University of Heidelberg, FRG 

MORZARIA, SUBHASH P., International Laboratory for Research on Animal Diseases, Kenya 

ROSALES, JOSE Luis E., Centre de Investigacion y de Estudios Avanzados del IPN, Mexico 

SAMARAS, NICHOLAS, Walter and Eliza Hall Institute of Medical Research, Australia 

SHONEKAN, OPEOLU A., University of Ibadan, Nigeria 

SINNIS, PHOTINI, Dartmouth Medical School 

STUCKY, PAMELA D., University of California, San Francisco 

TALAMAS, PATRICIA R., Centro de Investigacion y de Estudios Avanzados del IPN, Mexico 

WEIDANZ, WILLIAM P., Hahnemann University School of Medicine 

ZIMMERMAN, RONALD J., Vanderbilt University 

EMBRYOLOGY: A MODERN COURSE IN DEVELOPMENTAL BIOLOGY 

Course directors 

BRANDHORST, BRUCE, McGill University, Canada 
JEFFERY, WILLIAM, University of Texas 



EDUCATIONAL PROGRAMS 57 



Other faculty, staff, and lecturers 

ARNOLD, JOHN M, University of Hawaii 

CHILDS, GEOFFREY, Albert Einstein College of Medicine 

CLARK, WALLIS, Bodega Marine Station 

COSTANTINI, FRANK, Columbia University 

DAVIDSON, ERIC, California Institute of Technology 

ELDON, ELIZABETH, M. D. Anderson Hospital 

EMERSON, CHARLES, University of Virginia 

GERHART, JOHN, University of California, Berkeley 

GIMLICH, ROBERT, University of California, Berkeley 

GOLSTEYN, ROY, University of Calgary, Canada 

GROSS, PAUL, Marine Biological Laboratory 

HAFNER, MATHIAS, German Cancer Research Center, FRG 

HILLE, MERRILL, University of Washington 

JAENISCH, RUDOLF, Massachusetts Institute of Technology 

JAFFE, LAURINDA, University of Connecticut Health Center 

JAFFE, LIONEL, Marine Biological Laboratory 

KADO, RAYMOND, Centre National Recherche Scientifique, France 

KEMPHUES, KENNETH, Cornell University 

KLEIN, WILLIAM, M. D. Anderson Hospital 

KLINE, D., University of Connecticut 

LEE, JAMES, California Institute of Technology 

MARZLUFF, WILLIAM, Florida State University 

MEEDEL, THOMAS H., Marine Biological Laboratory 

RICHTER, JOEL, Worcester Foundation for Experimental Biology 

ROBERTS, JAMES, Hutchinson Cancer Center 

ROSBASH, MICHAEL, Brandeis University 

RUDERMAN, JOAN, Duke University 

SCHATTEN, GERALD, University of Wisconsin 

SCHATTEN, HEIDI, Florida State University 

SLUDER, GREENFIELD, Worcester Foundation for Experimental Biology 

WESSEL, GARY, M. D. Anderson Hospital 

WHITTAKER, J. RICHARD, Marine Biological Laboratory 

WILT, FRED, University of California, Berkeley 

WINKLER, MATTHEW, University of Texas, Austin 

WORMINGTON, MICHAEL, Brandeis University 

Students 

ANDERSON, MARYDILYS S., Yale University 

BERG, CELESTE A., Carnegie Institution/Yale University 

BICKEL, SHARON E., Baylor College of Medicine 

BLOOM, THEODORA L., University of Cambridge, England, UK 

BURSDAL, CAROL A., Duke University 

FORRESTER, WILLIAM C, University of Washington 

HAFNER, MATHIAS**, German Cancer Research Center, FRG 

HARDIN, PAUL E., Indiana University 

HARDIN, SUSAN H.. Indiana University 

HOULISTON, EVELYN, University of Cambridge, England, UK 

JURSNICH, VICTORIA A., University of California, Irvine 

KIRBY, COLLEEN M., Cornell University 

KOENIG, GERD, Max Planck Institut fur Entwicklungsbiologie, FRG 

KUBIAK, JACEKZ., Warsaw University, Poland 

RUBACHA, ALICE, Rice University 

** Advanced Research Training Program participant. 



58 MARINE BIOLOGICAL LABORATORY 

SAAVEDRA, CAROL, McGill University, Canada 

SCHOLER, ANNE-MARIE, Han/ard University 

SCHROETER, SALLY J., University of Michigan 

SMOLICH, BEVERLY D., University of Virginia 

SYMES, KAREN, National Institute of Medical Research, England, UK 

TALEVI, RICCARDO, University of Naples, Italy 

VARNUM, SUSAN M., Brandeis University 

VELLECA, MARK A., Yale University 

VITES, ANA M., University of Connecticut Health Center 

WHARTON, L. LYNN, University of Massachusetts Medical School 



MARINE ECOLOGY 
Course director 

FRANK, PETER W., University of Oregon 
Other faculty, staff, and lecturers 

ANDERSON, DONALD M., Woods Hole Oceanographic Institution 

Buss, LEO, Yale University 

CAPUZZO, JUDITH, Woods Hole Oceanographic Institution 

CARACO, NINA, Mary Flagler Cary Arboretum 

CARLTON, JAMES, Williams College 

CARON, DAVID A., Woods Hole Oceanographic Institution 

CASWELL, HAL, Woods Hole Oceanographic Institution 

CAVANAUGH, COLLEEN, Harvard University 

COLE, JON, Mary Flagler Cary Arboretum 

DAVIS, CABELL, Woods Hole Oceanographic Institution 

DELANO, M., Environmental Protection Agency 

DEUSER, WERNER G., Woods Hole Oceanographic Institution 

FOREMAN, KENNETH, Marine Biological Laboratory/BUMP 

FREADMAN, MARVIN, Marine Biological Laboratory/BUMP 

FROST, BRUCE W., University of Washington 

GALLAGHER, EUGENE, University of Massachusetts 

GAINES, ARTHUR G., JR., Woods Hole Oceanographic Institution 

GIBLIN, ANN, Marine Biological Laboratory 

GRASSLE, J. FREDERICK, Woods Hole Oceanographic Institution 

GRASSLE, JUDITH, Marine Biological Laboratory 

HARBISON, G. RICHARD, Woods Hole Oceanograpic Institution 

HOBBIE, JOHN E., Marine Biological Laboratory 

HUSTON, MICHAEL, Oak Ridge National Laboratory 

JEFFERIES, ROBERT L., University of Toronto, Canada 

MANN, KENNETH H., Bedford Institute of Oceanography, Canada 

MARCY, MARIBEL, Smith College 

MURCHELLANO, ROBERT, National Marine Fisheries Service 

OSMAN, RICHARD, Academy of Natural Sciences of Philadelphia 

PASCUAL-DUNLAP, M. MERCEDES, Cornell University 

PETERSON, CHARLES HENRY, University of North Carolina 

PETERSON, BRUCE R., Marine Biological Laboratory 

PREGNALL, MARSHALL, University of Massachusetts 

REX, MICHAEL, University of Massachusetts 

RHOADS, DONALD, Yale University 

RICE, DONALD, Chesapeake Biological Laboratory 

RUBLEE, PARKE A., Whitman College 

SANDERS, HOWARD L., Woods Hole Oceanographic Institution 

SEBENS, KENNETH, Northeastern University 



EDUCATIONAL PROGRAMS 59 

SHELLEY, PETER, Conservation Law Foundation 

VALIELA, IVAN, Boston University 

WALLACE, GORDON, University of Massachusetts 

WEINBERG, JAMES R., Woods Hole Oceanographic Institution 

WELSCHMEYER, NICHOLAS, Harvard University 

WIEBE, PETER H., Woods Hole Oceanographic Institution 

Students 

BROWN, ANNE C., University of Oregon 

COMIN, FRANCISCO A., University of Barcelona, Spain 

DICKENS, VIRGINIA A., Goucher College 

DIOGENE, GEORGES F., University of Barcelona, Spain 

DUBILIER, NICOLE, University of Hamburg, FRG 

FALK, KATHLEEN, University of Massachusetts, Boston 

FREY, IRIS J. F., Philipps-University Marburg, FRG 

HART, ROBERTA., University of California, Berkeley 

KASMER, JOHN M., University of Vermont 

MORUCCI, CARLO, University La Sapienza of Rome, Italy 

MYERS, PHILIP E., University of South Carolina 

O'HARA, ELLEN MARGARET, Villanova University 

SPANO, ANNAMARIA, Istituto Superiore de Sanita of Rome, Italy 

SVENDSEN, BETTY-ANN E., University of Dallas 

THIVAKARAN, ALAGIRI G., Annamalai University, India 

THOMAS, CECELIA R., Hinds Jr. College 

ZAPATA, FERNANDO A., University of Arizona 

MICROBIOLOGY: MOLECULAR ASPECTS OF CELLULAR DIVERSITY 
Course directors 

GREENBERG, PETER, Cornell University 
WOLFE, RALPH, University of Illinois 

Other faculty, staff, and lecturers 

ARMITAGE, JUDITH, Oxford University, UK 

BLAKEMORE, RICHARD, University of New Hampshire 

BOBIK, THOMAS, University of Illinois 

DILLING, WALTRAUD, University of Konstanz, FRG 

DiMARCO, ANTHONY, University of Illinois 

DUNLAP, PAUL, Cornell University 

FRANKEL, RICHARD, Massachusetts Institute of Technology 

JEFFERYS, JUDITH, Oxford University, UK 

KAPLAN, SAMUEL, University of Illinois 

KAISER, DALE, Stanford University 

KROPINSKI, ADAM, Marine Biological Laboratory 

KROPINSKI, ANDREW, Queen's University, Canada 

MACNAB, ROBERT, Yale University 

PFENNIG, NORBERT, Universitat Konstauz, FRG 

ROUVIERE, PIERRE, University of Illinois 

SPUDICH, JOHN, Albert Einstein College of Medicine 

WIDDEL, FRIEDRICH, University of Illinois, Urbana-Champaign 

WRAIGHT, COLIN A., University of Illinois 

Students 

ANDERSON, KAREN L., University of Iowa 
DOBBS, FREDC, Florida State University 



60 MARINE BIOLOGICAL LABORATORY 

GALLO, MARK A., Cornell University 
GIBSON, SUSAN A., University of Oklahoma 
KHANDEKAR, SANJAY S., Portland State University 
KING, STAGG L., U< isity of Washington 
KOT, MARK, ? . uty of Arizona 

KUTZ, Sus ' Diversity of Arizona 

LANE, DA. .. Indiana University 
Liu, SHU M., University of Oregon 
MANGIN. KATRINA L., University of Arizona 
MARCIKJK, DOUGLAS A., Hope College 
MICHEL, TOMAST., University of California, Davis 
PADGITT, PATRICIA J., Creighton University 
SILVERSTONE, SARA E., University of California, Davis 
SPORMANN, ALFRED MICHAEL, Philipps Universitat, FRG 
STEPHENS, CRAIG MICHAEL, University of Virginia 
STODDARD, STEVEN F., University of Wisconsin, Madison 
WEISS, DAVIDS., University of California, Berkeley 
ZHAO, HONGXUE, University of Illinois 



NEURAL SYSTEMS AND BEHAVIOR 

Course directors 

CAREW, THOMAS, Yale University 
KELLEY, DARCY, Columbia University 

Other faculty, staff, and lecturers 

AVITABLE, ELENA, Columbia University 

BASS, ANDREW, Cornell University 

BORST, AXEL, Max Planck Institut fur Cell Biologic, FRG 

BURD, GAIL, University of Arizona 

BYRNE, JOHN, University of Texas Medical School 

CALABRESE, RONALD, Harvard University 

CARROLL, LESLIE, Thomas Jefferson University 

CASAGRANDE, VIVIAN, Vanderbilt University 

CLEARY, LEONARD, University of Texas 

ELLIOT, ELLEN, University of North Carolina 

FRANK, JILLIAN, New York University 

GOLDMAN-RAKJC, PATRICIA, Yale University Medical School 

GORLICK, DENNIS, Columbia University 

HARRIS- WARRICK, RONALD, Cornell University 

HOSKINS, SALLY, Columbia University 

JACOBS, GWEN, University of California, Berkeley 

JOHNSON, BRUCE, Cornell University 

LEVINE, RICHARD, Rice University 

MACAGNO, EDUARDO, Columbia University 

MARDER, EVE, Brandeis University 

McROBERT, SCOTT, Temple University 

MOISEFF, ANDREW, University of Connecticut 

NORTHCUTT, GLENN, University of Michigan 

NUSBAUM, MICHAEL, Brandeis University 

PEARSON, KEIR, University of Alberta 

SIMMONS, JAMES, Brown University 

SQUIRE, LARRY, University of California, San Diego 

TOBIAS, MARTHA, Columbia University 



EDUCATIONAL PROGRAMS 

TOMPKINS, LAURIE, Temple University 
WEEKS, JANIS, University of California, Berkeley 

Students 

APPLEGATE, APRIL V., Johns Hopkins School of Medicine 

GLOWER, ROBERT P., Cornell University 

DODD, FRANK, Cornell University 

EDMONDS, BRIAN W., University of Virginia 

ELIOT, LISE SUZANNE, Columbia University 

GARCIA CABRERA, INMACULADA, University of Bergen, Norway 

HAMMER, MARTIN, Freie Universitat Berlin, FRG 

HARRINGTON, MARY E., Dalhousie University, Canada 

HIRANO, ARLENE A., The Rockefeller University 

KIEHN, OLE, The Panum Institute, Denmark 

KNOWLTON, BARBARA, Stanford University 

LoTuRCO, JOSEPH J., Yale University 

LUSTIG, CORNEL, Weizmann Institute, Israel 

MENCIO, TRACEY L., Rutgers University 

MORGAN, MICHAEL M., University of California, Los Angeles 

NIRENBERG, SHEILA, Harvard Medical School 

NISSANOV, JONATHAN, University of Colorado 

ROBERTS, SETH D., University of California, Berkeley 

STREICHERT, LAURA D., Stanford University 

WEAVER, DEBORAJ., University of Maryland, Baltimore County 



NEUROBIOLOGY 

Course director 

KARLIN, ARTHUR, Columbia University 

Other faculty, staff, and lecturers 

ADAMS, PAUL, SUNY, Stony Brook 

AGNEW, WILLIAM, Yale University 

ANDERSON, DAVID, Columbia University 

ANDREWS, BRIAN, NINCDS/NIH 

ARMSTRONG, KATIE, Rice University 

BRETT, ROGER, SUNY, Stony Brook 

CLAUDIO, TONI, Yale University Medical School 

CONNER, JOHN, Bell Laboratories 

DiPAOLA, MARIO, Columbia University College of Physicians and Surgeons 

EHRLICH, BARBARA, Albert Einstein College of Medicine 

FISCHBACH, GERALD, Washington University School of Medicine 

GURNEY, ALISON, California Institute of Technology 

HALL, LINDA, Albert Einstein College of Medicine 

HATTEN, MARY E., New York University Medical Center 

HESS, PETER, Harvard University 

HEUSER, JOHN, Washington University 

INOUE, TOMO, McGill University, Canada 

JESSELL, THOMAS, Columbia University 

JONES, STEVEN, SUNY, Stony Brook 

KHAN, SAHID, Marine Biological Laboratory 

LANDER, ARTHUR, Columbia University 

LANDIS, DENNIS, Massachusetts General Hospital 

LESTER, HENRY, California Institute of Technology 



62 MARINE BIOLOGICAL LABORATORY 

LEVITAN, IRWIN, Brandeis University 

MACKINNON, RODERICK, Brandeis University 

MARGULIES, DAVID, Columbia University College of Physicians and Surgeons 

MATSUMOTO, STEV , ; , Harvard University 

McNiVEN, !V ! J niversity of Maryland 

MILLER, O ;R, Brandeis University 

MOOSEK.E- , MARK, Yale University 

MURR . University of Pennsylvania 

PAULSEN, HENRY, Yale University Medical School 

RA VIOLA, ELIO, Harvard Medical School 

REFSE, THOMAS S., NINCDS/NIH/Marine Biological Laboratory 

ROLE, LORNA, Columbia University 

ROSENBLUTH, JOHN, New York University 

ROWLAND, L., Columbia University 

SCHNAPP, BRUCE, Marine Biological Laboratory 

SHEETZ, MICHAEL, Washington University School of Medicine 

SILMAN, ISRAEL, Weizmann Institute of Science, Israel 

SPUDICH, JOHN, Albert Einstein College of Medicine 

STERNWEIS, PAUL, University of Texas Health Center 

WILLARD, ALAN L., University of North Carolina 

Students 

BLEY, KEITH R., Yale University 
CARLBERG, MATS, University of Lund, Sweden 
FEDEROFF, HOWARD J., Massachusetts General Hospital 
FERNANDEZ- VALLE, CRISTINA, University of Miami 
HALPERN, MARNIE E., Yale University 
HORRIGAN, FRANK T., Stanford University 
LEW, DANIEL J., The Rockefeller University 
PLEASURE, SAMUEL J., University of Pennsylvania 
PORTER, DEVRA, Vanderbilt University 
SCHWEIZER, FELIX E., Universitat Basel, Switzerland 
SUPATTAPONE, SuRACHAi, Johns Hopkins School of Medicine 
ZUMBROICH, THOMAS J., University of Oxford, Oxford, UK 

PHYSIOLOGY 

Course director 

GOLDMAN, ROBERT, Northwestern University 
Other faculty, staff, and lecturers 

ALBRECHT-BUEHLER, G., Northwestern University 
BECKERLE, MARY, University of North Carolina 
BENDER, WELCOME, Harvard University 
BLOOM, KERRY, University of North Carolina 
CHISHOLM, REX, Northwestern University 
DEROSIER, DAVID, Brandeis University 
DESSEV, GEORGE N., Northwestern University 
FILETI, LISA, Boston University 
FUKUI, YOSHIO, Osaka University, Japan 
GOLDMAN, ANNE, Northwestern University 
GOLDSTEIN, LARRY, Harvard University 
HAN, PETER, Earlham College 
HAY, ELIZABETH D., Harvard University 
HOLM, CONNIE, Harvard University 



EDUCATIONAL PROGRAMS 63 



HOPKINSON, SUSAN, Northwestern University Medical School 

HORVITZ, H. ROBERT, Massachusetts Institute of Technology 

HYAMS, JEREMY, University College, UK 

JONES, JONATHAN, Northwestern University 

KIEHART, DAN, Harvard University 

LEINWAND, LESLIE, Albert Einstein College of Medicine 

LINDBERG, UNO, University of Stockholm, Sweden 

MAYRAND, SANDRA, Worcester Foundation for Experimental Biology 

MATTOX, ANDREW, Marine Biological Laboratory 

MCNALLY, ELIZABETH, Albert Einstein College of Medicine 

MORELAND, ROBERT, Dana Farber Cancer Institute 

OLMSTED, JOANNA, University of Rochester 

PARYSEK, LINDA, Northwestern University Medical School 

PEDERSON, THORLJ, Worcester Foundation for Experimental Biology 

PTASHNE, MARK, Harvard University 

RICH, ALEXANDER, Massachusetts Institute of Technology 

RUDERMAN, JOAN, Duke University 

RUSHFORTH, ALICE, Earlham College 

RUSKIN, BARBARA, Harvard University 

SCHWARTZ, LAWRENCE M., University of North Carolina 

SHAPIRO, LUCY, Albert Einstein College of Medicine 

SINGER, REBECCA, Albert Einstein College of Medicine 

SOHN, REGINA LEE, Albert Einstein College of Medicine 

SPUDICH, JAMES, Stanford University 

STEVENSON, BRUCE, Yale University 

SZENT-GYORGYI, ANDREW, Brandeis University 

TARDIFF, JILL, Albert Einstein College of Medicine 

TAYLOR, MARK, Northwestern University Medical School 

VALE, RON, Marine Biological Laboratory 

VALLEE, RICHARD, Worcester Foundation for Experimental Biology 

WARNER, CECELIA, Northwestern University Medical School 

WARNER, JONATHAN, Albert Einstein College of Medicine 

WEINSTEIN, RONALD, Rush Medical Center 

WHITMAN, GEORGE, Worcester Foundation for Experimental Biology 

WIEBEN, ERIC D., Mayo Foundation 

YEH, ELAINE, CIBA-GEIGY 

Students 

AKINS, JR., ROBERT E., University of Pennsylvania 
BEEMAN, ANNE M., Dartmouth Medical School 
BISWAS, SURAJIT K., University of Pennsylvania 
BLACK, KRISTIN, University of California, Berkeley 
BRADLEY, DAVID, University of Pennsylvania 
CAULEY, KEITH A., University of Michigan 
DABORA, SANDRA L., University of Connecticut 
DAHL, STEPHEN C, Wesleyan University 
DASSO, MARY C., Cambridge University, England, UK 
DEYST, KATHERINE A., Tufts University 
ERIKSSON, ULF J., Uppsala University, Sweden 
FOLTZ, KATHLEEN R., Purdue University 
GANNON, PAMELA M., Tufts University-Sackler School 
GELLES, JEFF, California Institute of Technology 
GUDEMAN, DAVID M., Kansas University 
HARPER, DAVIDS., University of Illinois, Chicago 
HORNE, MARY C., University of California, San Francisco 
KATZ, KENNETH S., Amherst College 



64 MARINE BIOLOGICAL LABORATORY 

KENNA, MARGARET A.. University of North Carolina 

KENNEY, LINDA!., University of Pennsylvania 

LAUERMAN, TOD V ' ' .>hns Hopkins University 

MEINHOF, C/ 3, University of California, San Diego 

MELUH, P\\fi University of Maryland 

MOHL, ViP , Washington State University 

REGINATO. ANTONIO M., University of Pennsylvania 

RUDOLFS KAREN M., Dartmouth College 

SARDET, CLAUDE C. S., Centre National de la Recherche Scientifique, France 

SEGRK. GINO V., Massachusetts General Hospital/Harvard Medical School 

SYMONS, MARC H. C., Weizmann Institute of Science, Israel 

THALER, CATHERINE D., University of California, Riverside 

TROXELL, CYNTHIA L., University of Colorado 

WADSWORTH, WILLIAM G., University of Missouri 

WATSON, CORNELIUS A., Wesleyan University 

YANAGIHARA, RICHARD, National Institute of Neurological and Communicative Disorders 

and Strokes/NIH 

YORK, KAREN PICKWICK, University of Pennsylvania 

ZAND, MARTIN S., Northwestern University Medical School 



SPRING 

ANALYTICAL AND QUANTITATIVE LIGHT MICROSCOPY IN BIOLOGY, 
MEDICINE, AND MATERIALS SCIENCE 

April 3- 10, 1986 
Course director 

INOUE, SHINYA, Marine Biological Laboratory 
Other faculty, staff, and lecturers 

AKINS, ROBERT, University of Pennsylvania 
ELLIS, GORDON W., University of Pennsylvania 
KNUDSON, ROBERTA., Marine Biological Laboratory 
LANNI, FREDERICK, Carnegie Mellon University 
LUBY-PHELPS, KATHERINE, Carnegie Mellon University 
LUTZ, DOUGLAS, Harvard University 
SALMON, EDWARD A., University of North Carolina 
TAYLOR, D. LANSING, Carnegie Mellon University 
WALKER, RICHARD, University of North Carolina 

Commercial faculty 

ABROMOWITZ, MORTIMER, Olympus Corporation of America 

ALEXANDER, SCOTT, Nikon, Inc. 

BEACH, DAN, Carl Zeiss, Inc. 

BREEN, BILL, Interactive Video Systems 

DEMIAN, JEFFREY, Nikon, Inc. 

ESSER, HERMAN, Ikegami Electronics (USA), Inc. 

FOSTER, BARBARA, Carl Zeiss, Inc. 

GRACE, JOHN, Crimson Camera Technical Sales, Inc. 

HANNAWAY, WYNDHAM, G. W. Hannaway Associates 

HINSCH, JAN, E. Leitz, Inc. 

JONES, JEFFREY, Olympus Corporation of America 

KELLER, ERNST, Carl Zeiss, Inc. 

KIMURA, T., Olympus Corporation of America 



EDUCATIONAL PROGRAMS 65 

KLEIFGEN, GERRY, Dage-MIT 

KNUTRUD, PAUL, Interactive Video Systems 

ORWELL, PATTY, E. Leitz, Inc. 

PACKARD, MEL, Quantex Corporation 

PRESLEY, PHIL, Carl Zeiss, Inc. 

REGAN, ANN, Ikegami Electronics (USA), Inc. 

RUBINOW, JERRY, Universal Imaging Corporation 

SCHEIRER, KURT, Nikon Inc. 

SCOTT, ERIC, Ikegami Electronics (USA), Inc. 

TAYLOR, RICHARD, Colorado Video 

THOMAS, PAUL, Dage-MIT 

VRATNEY, MELANA, Nikon Inc. 

Students 

ANDERSON, DONALD M., Woods Hole Oceanographic Institution 

ARONSON, JOHN F., Wistar Institute 

BARI, DANIEL, Universidad Nacional de Cuyo-Conicet, Argentina 

CAVICCHIA, JUAN CARLOS, Universidad Nacional de Cuyo-Conicet, Argentina 

CHAMBERS, EDWARD L., University of Miami School of Medicine 

DUBINSKY, JANET M., Washington University School of Medicine 

FEIGENSON, GERALD W., Cornell University 

GALLAGER, SCOTT M., Woods Hole Oceanographic Institution 

GLANZMAN, DAVIDL., Howard Hughes Medical Institute 

KHAN, SHAHID M. M., Albert Einstein College of Medicine 

KLEITMAN, NAOMI, Washington University School of Medicine 

McCuLLOH, DAVID H., University of Miami School of Medicine 

MILLER, PAUL, Bell Laboratories 

NORRIS, CAROLYN, Bardeen Labs 

PALAZZO, ROBERT E., University of Virginia 

REESE, TOM, Marine Biological Laboratory 

RUSSELL, JAMES T.,NIH 

SCHULZE, ERIC, University of California, San Francisco 

SHEEHY, PAULA., NIH 

VALE, RONALD D., Marine Biological Laboratory 

VOYVODIC, JAMES, Washington University School of Medicine 

WIGSTON, DONALD, Emory University School of Medicine 

WOMACK, MARY, Howard Hughes Medical Institute 

YEAGER, MARK D., Cornell University 



SHORT COURSES 

CELL AND MOLECULAR BIOLOGY OF PLANTS 

August 4- 14, 1986 
Coordinators 

DURE, LEON, The University of Georgia 
KEY, JOE L., The University of Georgia 

Lecturers 

AUSUBEL, FRED, Massachusetts General Hospital 

CHUA, NAM, Rockefeller University 

CLEGG, MICHAEL, University of California, Riverside 

CROUCH, MARTHA, Indiana University 

FRALEY, ROB, Monsanto Company 



66 MARINE BIOLOGICAL LABORATORY 

HEPLER, PETER K., University of Massachusetts 
LEVINGS, C. S. III. North Carolina State University 
MALMBERG, RUSSEI.L, The University of Georgia 
MEAGHER, RICHARD, The University of Georgia 
PALEVITZ, BARRY, The University of Georgia 
QUAIL, PFTI R, University of Wisconsin 
SOMMe : ; v LE, CHRIS, Michigan State University 
STROI. i, JUDITH, The University of Georgia 
TLM; /.E, BILL, University of California, Davis 
VARNER, JOE, Washington University 
YODER, OLIN, Cornell University 

Students 

AGARWAL, MUNNA LAI, Centre National de la Recherche Agronomiue, France 

ARMOUR, SUSAN, CIBA-GEIGY Corp. 

ARMOUR, TOBY, Edgartown, Massachusetts 

BECK, JAMES J., CIBA-GEIGY Corp. 

CAROZZI, NADINE, CIBA-GEIGY Corp. 

CHENEY, DONALD P., Northeastern University 

CURRY, L. JEANNE, University of Massachusetts 

ELLIOTT, WILLIAM, Hartwick College 

GOLDMAN, PEG, New Haven, Connecticut 

HANLEY, SUSAN, BioTechnical International Inc. 

HAUGE, BRIAN, Massachusetts General Hospital 

LOTSTEIN, RICHARD, CIBA-GEIGY Corp. 

McCABE, BRIAN, Bloomington, Indiana 

MINEO, LORRAINE, Lafayette College 

MULCARE, DONALD J., Southeastern Massachusetts University 

NAM, HONG GIL, Massachusetts General Hospital 

NOBLE, REGINALD D., Bowling Green State University 

ROSE, VIRGINIA, Concord, Massachusetts 

SAXENA, INDER MOHAN, University of Texas, Austin 

WILLIAMS, SHERICCA C., CIBA-GEIGY Corp. 

BASIC IMMUNOCYTOCHEMICAL TECHNIQUES IN 
TISSUE SECTIONS AND WHOLE MOUNTS 

October 19-25, 1986 
Course directors 

BELTZ, BARBARA, Harvard Medical School 
BURD, GAIL D., University of Arizona 

Course assistants 

KENT, CARLA, University of Arizona 
KOBIERSKI, LINDA, Harvard Medical School 

Students 

AKANA, SUSAN FONG, University of California, San Francisco 
BRADFUEHRER, PETER D., Cornell University 
FOERSTER, ANNE, McMaster University, Canada 
GETCHELL, MARILYN L., Wayne State University 
HAMILTON, KATHRYN A., New England Medical Center 
HAMMAR, KATHERINE M., Marine Biological Laboratory/NIH 
HELLUY, SIMONE, The University of Alberta, Canada 



EDUCATIONAL PROGRAMS 67 



KRUSZEWSKA, BARBARA, University of Texas, Austin 

LAUFER, HANS, University of Connecticut 

Ross, LINDA S., University of Texas, Austin 

SASAVAGE, NANCY L., Bethesda Research Laboratories 

TiLSON, HUGH A., National Institute of Environmental Health Sciences 

WHITE, JOEL, The Florida State University 

WOOD, SUSAN F., Marine Biological Laboratory/BUMP 

ZIGMOND, RICHARD E., Harvard Medical School 



X. RESEARCH AND TRAINING PROGRAMS 

SUMMER 

PRINCIPAL INVESTIGATORS 

ALLEN, NINA S., Wake Forest University 

ALLEN, LABORATORY, Dartmouth College 

ANDERSON, WINSTON A., Hunter College 

ARMSTRONG, CLAY M., University of Pennsylvania 

ARMSTRONG, PETER B., University of California, Davis 

ATWOOD, KIM, Marine Biological Laboratory 

AUGUSTINE, GEORGE, University of Southern California 

BARRY, DANIEL, University of Michigan 

BARLOW, ROBERT B., Syracuse University 

BARRY, M., Albert Einstein College of Medicine 

BARRY, SUSAN R., University of Michigan 

BEAUGE, Luis ALBERTO, Institute de Investigacion Medica, Argentina 

BEGENISICH, TED, University of Rochester Medical Center 

BENNETT, MICHAEL V. L., Albert Einstein College of Medicine 

BEZANILLA, FRANCISCO, University of California, Los Angeles 

BLUNDON, JAY A., University of Maryland 

BODZNICK, DAVID, Wesleyan University 

BORGESE, THOMAS A., Lehman College 

BORON, WALTER F., Yale University 

BOYER, BARBARA C, Union College 

BRADY, SCOTT T., University of Texas Health Science Center 

BREHM, PAUL, Tufts University School of Medicine 

BROWN, JOEL E., Washington University 

BURDICK, CAROLYN J., Brooklyn College 

BURGER, MAX M., University of Basel, Switzerland 

CARMELIET, PETER, University of Leuven, Belgium 

CHANG, DONALD C., Baylor College of Medicine 

CHAPPELL, RICHARD L., Hunter College 

CHARLTON, MILTON P., University of Toronto, Canada 

CLEVELAND, MARK V. B., Braintree Laboratories 

COHEN, LAWRENCE B., Yale University 

COHEN, WILLIAM D., Hunter College 

CONDOURIS, GEORGE A., New Jersey Medical School 

COOPERSTEIN, SHERWIN J., University of Connecticut 

CORNWALL, M. CARTER, Boston University School of Medicine 

D'AVANZO, CHARLENE, Hampshire College 

DEWEER, PAUL, Washington University 

DUBAS, FRANCOISE, Whitney Laboratory of Marine Biology 

DUBE, FRANCOIS, University of Quebec 

DUNLAP, KATHLEEN, Tufts University School of Medicine 



68 MARINE BIOLOGICAL LABORATORY 

ECKBERG, WILLIAM R.. Howard University 

EHRLICH, BARBARA, Albert Einstein College 

FEINMAN, RICHARD, SUN Y Health Sciences Center 

FESTOFF, BARRY W.. University of Kansas Medical Center 

FISHMAN, HARVFV M., University of Texas Medical Branch 

FREADMAN, MARVIN, Marine Biological Laboratory 

FULKERSON, JOHN PRYOR, University of Connecticut School of Medicine 

GADSBY. DAVIDC., Rockefeller University 

GAINER, HAROLD, NICHD/NIH 

GARBER, SARAH S., Brandeis University 

GEORGE, EDWIN B., Case Western Reserve University 

GILBERT, DANIEL L., NINCDS/NIH 

GIUDITTA, ANTONIO, University of Naples, Italy 

GOULD, ROBERT, New York Institute for Basic Research 

GOVIND, C. K., University of Toronto 

GRAF, WERNER M., Rockefeller University 

HALVORSON, HARLYN O., Brandeis University 

HEPLER, PETER K., University of Massachusetts 

HIGHSTEIN, STEPHEN M., Washington University 

HILL, ROBERT B., University of Rhode Island 

HILL, SUSAN DOUGLAS, Michigan State University 

HOSKIN, FRANCIS C. G., Illinois Institute of Technology 

HORN, RICHARD, University of California Medical School 

HUMPHREYS, TOM, University of Hawaii 

JOHNSON, KENNETH A., Pennsylvania State University 

JOSEPHSON, ROBERT K., University of California, Irvine 

KALTENBACH, JANE C., Mount Holyoke College 

KAMINER, BENJAMIN, Boston University 

KAO, PETER, Columbia University 

KAPLAN, EHUD, Rockefeller University 

KEYNAN, ALEXANDER, Memorial Sloan Kettering Cancer Center 

KEM, WILLIAM R., University of Florida 

KORNBERG, HANS, University of Cambridge, UK 

LANDOWNE, DAVID, University of Miami 

LANGFORD, GEORGE M., University of North Carolina 

LASER, RAYMOND J., Case Western Reserve University 

LAUFER, HANS, University of Connecticut 

LEVIS, RICHARD A., Rush Medical Center 

LINDGREN, CLARK, Duke University Medical Center 

LIPICKY, RAYMOND JOHN, Food and Drug Administration 

LISMAN, JOHN, Brandeis University 

LLINAS, RUDOLFO R., New York University 

LOEWENSTEIN, WERNER R., University of Miami 

MALBON, CRAIG C., State University of New York, Stony Brook 

MATTESON, DONALD R., University of Pennsylvania 

METUZALS, J., University of Ottawa, Canada 

MORRELL, FRANK, Rush-Presbyterian-St. Luke's Medical Center 

MORRELL, LEYLA DE TOLEDO, Rush-Presbyterian-St. Luke's Medical Center 

MULLINS, LORIN J., University of Maryland 

NARAHASHI, TOSHIO, Northwestern University 

NELSON, LEONARD, Medical College of Ohio 

NOE, BRYAN D., Emory University 

NOLEN, THOMAS G., Yale University 

OHKI, SHINPEI, State University of New York, Buffalo 

PALEVITZ, BARRY A., University of Georgia 



RESEARCH AND TRAINING PROGRAMS 69 

PARSONS, THOMAS D., University of Pennsylvania School of Dental Medicine 

PALMER, JOHN D., University of Massachusetts, Amherst 

PEROZO, EDUARDO, Institute Venezolano de Investigaciones Cientificas, Venezuela 

PIERSON, BEVERLY K., University of Puget Sound 

POOLE, THOMAS J., Upstate Medical Center 

PUMPLIN, DAVID W., University of Maryland 

PURVES, DALE, Washington University 

QUIGLEY, JAMES P., SUNY, Downstate Medical Center 

RAKOWSKI, ROBERT F., Chicago Medical School 

REBHUN, LIONEL I., University of Virginia 

REID, JOHN, Hampshire College 

RENDER, JOANN, Hamilton College 

REYNOLDS, GEORGE T., Princeton University 

RICKLES, FREDERICK R., University of Connecticut Health Center 

RIPPS, HARRIS, University of Illinois College of Medicine 

ROME, LAWRENCE C, University of Tennessee 

Ross, WILLIAM, New York Medical College 

RONAN, MARK, Wesleyan University 

RUDERMAN, JOAN V., Duke University 

RUSSELL, JOHN M., University of Texas 

SALZBERG, BRIAN M., University of Pennsylvania 

SANGER, JOSEPH W., University of Pennsylvania 

SARDET, CHRISTIAN, Station Zoologique, France 

SCHROER, TRINA, University of California, San Francisco 

SEGAL, SHELDON J., Rockefeller Foundation 

SILVER, ROBERT B., University of Wisconsin 

SJODIN, RAYMOND A., University of Maryland School of Medicine 

SLOBODA, ROGER D., Dartmouth College 

SMITH, STEPHEN J., Yale University School of Medicine 

SPECK, WILLIAM T., Rainbow Babies & Childrens Hospital 

SPIEGEL, EVELYN, Dartmouth College 

SPIEGEL, MELVIN, Dartmouth College 

STANLEY, ELisF., NINCDS/NIH 

STEPHENS, PHILIP J., Villanova University 

STOCKBRIDGE, NORMAN, University of Alberta, Canada 

STRACHER, ALFRED, SUNY Health Sciences Center 

STRUMWASSER, FELIX, Boston University 

STUART, ANN E., University of North Carolina 

TAKEDA, KENNETH, Universite Louis Pasteur, France 

TASHIRO, JAY SHIRO, Kenyon College 

TAYLOR, ROBERT E., NINCDS/NIH 

TILNEY, LEWIS, University of Pennsylvania 

TRAVIS, JEFFREY L., Vassar College 

TREISTMAN, STEVEN N., Worcester Foundation 

TRINKAUS, JOHN PHILIP, Yale University 

TROLL, WALTER, New York University 

TUCKER, EDWARD B., Vassar College 

VINCENT, WALTER S., University of Delaware 

WAITE, MOSELEY, Bowman Gray School of Medicine 

WEIDNER, EARL, Louisiana State University 

WEISS, DIETER G., Institute of Zoology, FRG 

WEISSMAN, GERALD, New York University Medical Center 

WHITE, ROY L., Albert Einstein College of Medicine 

YEH, JAY Z., Northwestern University 

ZIGMAN, SEYMOUR, University of Rochester School of Medicine 



70 MARINE BIOLOGICAL LABORATORY 

LIBRARY READERS 

ADELBERG, EDWARD, Yale Medical School 

AKINS, KATHLEEN. Tufts University 

ALKON, DANIEL, NIH/NINCDS 

ALLEN, GARLAND E., Washington University 

ANDERSON. EVERETT, Harvard Medical School 

APOSHIAN, H. VASKEN, University of Arizona 

BABITSKY, STEVEN, Kistin, Babitsky, Latimer & Beitman 

BANG, BETSY, MBL 

BARRETT, DENNIS, University of Denver 

BEAN, CHARLES, Rensselaer Polytechnic Institute 

BEMIS, WILLY, University of Massachusetts 

BOETTIGER, JULIE, Temple University 

BOYER, JOHN, Union College 

BROWNE, ROBERTA., Wake Forest University 

BUCK, JOHN, NIH 

BURSZTAJN, S., Baylor College of Medicine 

CANDELAS, GRACIELA C., University of Puerto Rico 

CARRIERS, RITA, Downstate Medical Center 

CASAGRANDE, VIVIEN A., Vanderbilt University 

CHAMBERS, EDWARD L., University of Miami School of Medicine 

CHEN, CHONG, Boston University Marine Program 

CHILD, FRANK, Trinity College 

CLARK, ARNOLD, MBL 

COBB, JEWEL PLUMME, California State University 

COHEN, LEONARD A., American Health Foundation 

COHEN, SEYMOUR S., MBL 

COLEN, B. D., Newsday 

D'ALESSIO, GIUSEPPE, University of Naples 

DETTBARN, WOLF-D., Vanderbilt University 

DIPEOLU, OLUSEGUN O., Tuskegee University 

DUNCAN, THOMAS K., Nichols College 

EBERT, JAMES D., Carnegie Institution of Washington 

ECKBERG, WILLIAM D., Howard University 

ELLNER, JEROLD, Case Western Reserve University 

FARB, DAVID, SUNY 

FARMANFARMIAN, A., Rutgers University 

FEINGOLD, DAVID S., New England Medical Center 

FELDMAN, SUSAN, New Jersey Medical School 

FIELD, GEORGE, Center for Astrophysics 

FISHER, SAUL H., Milhauser Laboratory 

FRIENKEL, KRYSTYNA, NYU Medical Center 

FRIEDLER, GLADYS, Boston University School of Medicine 

FRENKEL, NORBERT, Northwestern University Medical School 

FUSSELL, CATHARINE P., Pennsylvania State University 

GERMAN, JAMES L., New York Blood Center 

GEWURZ, HENRY, Evanston, Illinois 

GOLDSTEIN, MOISE H., JR., Johns Hopkins University 

GOODGAL, SOLH., University of Pennsylvania School of Medicine 

GOUNARIS, ANNE D., Vassar College 

GRANT, PHILIP, University of Oregon 

GROSSMAN, ALBERT, NYU 

GUTTENPLAN, JOSEPH B., NYU Dental Center 

HARDING, CLIFFORD V., Kresge Eye Institute 

HAZEL, LEA, Kenyon College 

HERSKOVITS, THEODORE T., Fordham University 



RESEARCH AND TRAINING PROGRAMS 7 1 



HILDEBRAND, JOHN G., University of Arizona 

HILL, RICHARD W., Michigan State University 

HILLMAN, PETER, Hebrew University 

HILTS, PHILIP J., Washington Post 

ILAN, JOSEPH, Case Western Reserve University 

ILAN, JUDITH, Case Western Reserve University 

INOUE, SADAYUKI, McGill University 

JACOBS, LISA, Kenyon College 

KALAT, JAMES W., North Carolina State University 

KALTENBACH, JANE C, Mt. Holyoke College 

KARUSH, FRED, University of Pennsylvania 

KARUSH, WILLIAM, California State University 

KELLY, ROBERT, University of Chicago, College of Medicine 

KLEIN, DAVID, University of California, San Francisco 

KLEIN, MORTON, Temple University Medical School 

KLEMOW, KENNETH M., Wilkes College 

KRANE, STEPHEN M., Massachusetts General Hospital 

LADERMAN, AIMLEE, MBL 

LAZAROW, PAUL B., Rockefeller University 

LEE, JOHN J., City College of CUNY 

LEIGHTON, JOSEPH, The Medical College of Pennsylvania 

LEVITZ, MORTIMER, NYU Medical Center 

LLOYD, DAN, Simmons College 

LONG, CAROLE A., Hahnemann University School of Medicine 

LORAND, LASZLO, Northwestern University 

LYNCH, ELIZABETH, Kenyon College 

MACKENZIE, DEBORA OLIVIA, New Scientist 

MACLEISH, WILLIAM H., Houghton-Mifflin 

MAIENSCHEIN, JANE, Arizona State University 

MARFEY, ANNE, Danish Writers Guild 

MARINE RESEARCH, INC. 

MASER, MORTON, Woods Hole Educational Associates 

MATSUMURA, FUMIO, Michigan State University 

MAUTNER, HENRY G., Tufts University School of Medicine 

MAUZERALL, DAVID, Rockefeller University 

MCCANN-COLLIER, MARJORY, St. Peter's College 

McCoY, FLOYD W., Lamont-Doherty Geological Observatory 

MELE, SUZANNAH, Kenyon College 

MILLER, JULIE ANN, Science News 

MILLER, MELISSA, Kenyon College 

MILLS, ERIC L., Dalhousie University 

MITCHELL, RALPH, Harvard University 

MIZELL, MERLE, Tulane University 

MONROY, ALBERTO, Naples Zoological Station 

MOORE, JOHN W., Duke University Medical Center 

MORSE, PATRICIA M., Northeastern University 

MUSACCHAI, X. J., University of Louisville 

NAGEL, RONALD L., Albert Einstein College of Medicine 

NICKERSON, PETER A., SUNY, Buffalo 

OLINS, ADA L., University of Tennessee, Oak Ridge 

OLINS, DONALD E., University of Tennessee, Oak Ridge 

OLSZOWKA, ALBERT J., SUNY, Buffalo 

OTT, KAREN, University of Evansville 

PEISACH, JACK, Albert Einstein College of Medicine 

PERSON, PHILIP, VA Medical Center, Brooklyn, New York 

POOLE, ALAN F., MBL 

PRICE, BILL, Kenyon College 



72 MARINE BIOLOGICAL LABORATORY 

PROVASOLI, LUIGI, Yale University 

PRUSCH, ROBERT, Gonzaga University 

RAEBURN, PAUL, Associated Press 

REINER, JOH^' M. Vibany Medical College 

RINGER, STEPHKV. Childrens Hospital 

ROBINSON VI BL 

ROTH, Ei 

RUSSELL, H., University of Arizona College of Medicine 

Ru HUNTER, W. D., Syracuse University 

SCMit SINGER, R. WALTER, University of Medicine and Dentistry of New Jersey 

SCHMIDT, SUSANNE, Cape Cod Planning and Economic Development 

SEAVER, GEORGE, Seaver Assoc. 

SHAPLEY, ROBERT, Rockefeller University 

SHEMIN, DAVID, Northwestern University 

SHEPARD, FRANK, Woods Hole Data Base 

SHEPRO, DAVID, Boston University 

SHRIFTMAN, MOLLIE STARR, North Nassau Mental Health Center 

SLUDER, GREENFIELD, Worcester Foundation for Experimental Biology 

SOHN, JOEL, Joel Sohn Seafood 

SPECTOR, ABRAHAM, Columbia University 

SPOTTE, STEPHEN, Mystic Marinelife Aquarium 

STEINBERG, MALCOLM S., Princeton University 

STEPHENS, MICHAEL J., Rutgers University 

STEPHENSON, WILLIAM K., Earlham College 

STEVENS, CHARLES F., Yale Medical School 

SZENT-GYORGYI, ANDREW G., Brandeis University 

SZENTKIRALYI-SZENT-GYORGYI, EVA M., Brandeis University 

TONE, JEFFERSON, Kenyon College 

TRACER, WILLIAM, Rockefeller University 

TUTTLE, FRANK, Kenyon College 

TWEEDEL, KENYON S., University of Notre Dame 

VAN HOLDE, K. E., Oregon State University 

WAGNER, ROBERT R., University of Virginia 

WAINIO, WALTER, Rutgers University 

WANGH, LAWRENCE, Brandeis University 

WARREN, LEONARD, Wistar Institute 

WEBB, H. MARGUERITE, MBL 

WEINER, JONATHAN, Doylestown, Pennsylvania 

WHEELER, GEORGE E., Brooklyn College 

WHITTENBERG, BEATRICE, Albert Einstein College of Medicine 

WHITTENBERG, JONATHAN, Albert Einstein College of Medicine 

WlCHTERMAN, RALPH, MBL 

WILBUR, CHARLES G., Colorado State University 

WOLKEN, JEROME J., Carnegie Mellon University 

WORGUL, BASIL V., College of Physicians and Surgeons, Columbia University 

YATHIRAJ, SANJAY, Kenyon College 

YOUNG, WISE, NYU Medical Center 

Yow, F. W., Kenyon College 

ZACKS, SUMNER I., The Miriam Hospital 

ZIMMERMAN, MORRIS, Merck Sharp & Dohme Research Laboratory 

ZOTTOLI, STEVEN J., Williams College 

OTHER RESEARCH PERSONNEL 

ABRAHAMIAN, LORI, University of Connecticut Health Center 
ABRAMSON, CHARLES, State University of New York Health Sciences Center 



RESEARCH AND TRAINING PROGRAMS 73 

ALEXANDER, R. MCNEILL, University of Leeds, UK 

ALTAMIRANO, ANIBAL, University of Texas 

ARMSTRONG, SANDRA, Lower Merion High School 

ASHLEY, C. C, Oxford University 

BAKER, ROBERT, New York University 

BAKER, Ross, University of Connecticut Medical Center 

BATES, HISLA, Hunter College 

BENNETT, ELENA P., Connecticut College 

BISIER, ENRIQUE FONT, University of Tennessee 

BLECK, THOMAS P., Rush Medical College 

BLUMER, JEFFREY L., Rainbow Babies and Children's Hospital 

BORST, DAVID, Illinois State University 

BOYLAN, JEANETTE, Michigan State University 

BREITWEISER, GERDA E., University of Texas Medical Branch 

BROSIUS, D., Albert Einstein College of Medicine 

BROWN, LESLEE DODD, Northwestern University Medical School 

BROWNE, CAROL, Wake Forest University 

BUTNAM, JOHN A., Washington University School of Medicine 

CAPUTO, CARLO, Institute Venezolano de Investigaciones Cient 

CARIELLO, Lucio, Naples Zoological Station, Italy 

CATTARELLI, MARTINE H., Yale University School of Medicine 

CALLAWAY, JAY, University of Washington 

CATANEO, RENE, NYS Institute for Basic Research in Developmental Disabilities 

CHANDLER, ROBERT, University of Maryland 

CHOW, ROBERT H., University of Pennsylvania 

CHEN, ERIC, Northwestern University 

CLARK, GEOFF, Braintree Laboratories 

COLTON, CAROL, Georgetown University Medical School 

COHEN, AVRUM, University of Chicago 

COTA-PENUELAS, GABRIEL, University of Pennsylvania 

COUCH, ERNEST, Texas Christian University 

COTE, RICK, University of Wisconsin 

CZINN, STEVEN, Rainbow Babies and Childrens Hospital 

DAVIDSON, DAVID, New York University Medical School 

DAVIDSON, SARAH, Columbia University 

DEWEILLE, JAN, University of Utrecht, Netherlands 

DIPOLO, REINALDO, Institute de Investigacion Medica, Argentina 

DIXON, ROBERT, College of the Holy Cross 

DOME, JEFF, University of Pennsylvania School of Medicine 

DOWLING, JOHN E., Harvard University 

DOUGHERTY, KATHLEEN, University of Delaware 

DUAX, J. B., SUNY, Buffalo 

DULDULAO, MARLYN, University of Hawaii 

EATON, D. C., University of Texas Medical Branch 

EHRENSTEIN, DAVID, Oberlin College 

EHRENSTEIN, G., NINCDS/NIH 

FINK, RACHEL D., Mount Holyoke College 

FLACKER, JONATHAN M., Emory University 

FONG, C. N., University of Toronto, Canada 

FRANK, DOROTHY, Rainbow Babies and Childrens Hospital 

GILBERT, SUSAN P., Dartmouth College 

GONSALVES, NEIL, Rhode Island College 

GONZALEZ, HUGO, University of Maryland 

GRAUBARD, KATHERINE, University of Washington 

GRAY, DAVID A., University of Southern California 

GREINER, FRANCINE, Emory University 



74 MARINE BIOLOGICAL LABORATORY 

GRIFFITHS, PETER J., University Laboratory of Physiology 

GRUNER, JOHN A., New York University 

GREEN, WENDY B., Amherst College 

HANTAI, DANIEL, 1 M.S.E.R.M. 

HEITHAUS, E, R., Kenyon College 

HERLANDS, Louis, Population Council 

HIRIART, MARCIA, University of Pennsylvania 

HOCSON. DANIAL, University of Puget Sound 

HOLBROOK, PAMELA G., Massachusetts Institute of Technology 

HOLDER, DAVID, City University of New York 

HOUGHTON, SUSAN, Marine Biological Laboratory 

HOMOLA, ELLEN, University of Connecticut 

HUNT, TIM, University of Cambridge, UK 

HUNT, JOHN R., Baylor College of Medicine 

IVENS, KEITH, Howard University 

JACKSON, LOVERNE, University of Ottawa, Canada 

JOCKUSCH, BRIGITTE M., University of Bielefeld, FRG 

JOHNSON, EDWIN, Brandeis University 

KASS, LEONARD, University of Maine 

KAHLER, CHERYL, Kansas City Veterans Administration Medical Center 

KAHN, TERRI, Rainbow Babies and Childrens Hospital 

KEM, ELAINE S., Fairleigh Dickinson University 

KEM, JAMES, University of Florida 

KISHIMOTO, YASUA, Johns Hopkins School of Medicine 

KNAKAL, ROGER C., Yale University 

KNIER, JULIE A., University of Minnesota 

KOIDE, SAMUEL S., Population Council 

KONZELMANN, DANIEL J., Eastern Illinois University 

KOSIK, K. S., Harvard University Medical School 

LANDOLFA, MICHAEL A., Union College 

LANDAU, MATTHEW, University of Connecticut 

LEECH, COLIN A., University of Cambridge, UK 

LEHMAN, HERMAN, Rockefeller University 

LEOPOLD, PHILIP LUTZ, Georgetown University 

LONDON, JILL, Yale University School of Medicine 

LOPEZ-BARNEO, JOSE, University of Seville Medical School, Spain 

LOPICCOLO, DANIEL, Medical College of Ohio 

LUCA, FRANK, Duke University Medical Center 

LUTZ, GORDON, University of Tennessee 

MACK, ERIN, University of Puget Sound 

MAMUYA, WILFRED, Boston University School of Medicine 

MASSEY, ERIC, University of North Carolina 

MASSIOTTE, J. MATHIEU, University of Connecticut Health Center 

MCCARTHY, ROBERT ALAN, University of Basel, Switzerland 

McGuiNNESS, T., Rockefeller University 

MELLO, ANIBEL, Rhode Island College 

MENICHINI, ENRICO, Northwestern University 

MERRITT, MARIA, Wake Forest University 

MEYER, MONICA A., Vassar College 

MILLER, ROBERT, Case Western Reserve University 

MILLS, VAN, The University of North Carolina 

MISEVIC, GRADIMIR, University of Basel, Switzerland 

MOCHEL, SUSAN, Tufts University 

MURRAY, SANDRA, University of Pittsburgh 

NAKA, KEN-!CHI, National Institute of Basic Biology, Japan 

NICHOLAS, CRAIG JOHN, Syracuse University 



RESEARCH AND TRAINING PROGRAMS 75 

NISHIO, MATOMO, Northwestern University Medical School 

OBAID, ANA LIA, University of Pennsylvania 

ORTIZ, ROSALEE, Howard University 

OSSES, Luis, University of California, Los Angeles 

PALAZZO, ROBERT, University of Virginia 

PANT, HARISH, NIAAA/NIMH/DHHS 

PAXHIA, TERESA M., University of Rochester 

PAXSON, CHERYL, University of Chicago Medical School 

PAULSEN, REINHARD, Ruhr University, FRG 

PEREZ, ROSA, Hunter College 

RALPH, WALTER, City University of New York 

RASGADO-FLORES, HECTOR, University of Maryland 

RENDER, TIMOTHY JOHN, University College, Oxford, UK 

REQUENA, JAIME, I.D.E.A., Venezuela 

RIESEN, WILLIAM J., Yale University 

ROSE, BIRGIT, University of Miami 

ROBINSON, JoHNT., University of North Carolina 

ROBINSON, PHYLLIS, Brandeis University 

ROSENBAUM, ROBERT, Vassar College 

RUDOLPH, REBECCA, University of Puget Sound 

SANDS, VICKJ, University of Puget Sound 

SANGER, JEAN, University of Pennsylvania 

SAHNI, MUKESH, Rockefeller Foundation 

SAK.AI, HIROKO, National Institute for Basic Biology, Japan 

SAWYER, PAM, University of Ottawa, Canada 

SCHLUP, VERENA, University of Basel, Switzerland 

SCHNEIDER, ERIC, Wesleyan University 

SCHIMINOVICH, DAVID, Yale University 

SCHNEIDER, MELISSA, Hamilton College 

SEITZ-TUTTER, DIETER, Institute fur Zoologie, FRG 

SHEETZ, JENNIFER, Duke University 

SHEN, JOANNE, University of Southern California 

SIEGAL, NINA, Case Western Reserve University 

SIMPSON, MARCIA, Amherst College 

SOLOMON, JOEL, Washington University School of Medicine 

SPIRES, SHERRILL, University of Rochester Medical Center 

STEINACKER, ANTOINETTE, Washington University School of Medicine 

STOCKBRIDGE, LISA, University of Alberta, Canada 

STOKES, DARRELL, Emory University 

STRONG, JOHN C., University of Maryland, Baltimore 

SUGIMORI, MUTSUYUKI, New York University 

SWANDULLA, DIETER, University of Pennsylvania 

SWENSON, KATHERINE, Harvard University School of Medicine 

TAKEDA, KIMIHISA, Tottori University, Japan 

TAKLA, NORA, Washington University School of Medicine 

TANGUY, JOELLE, Laboratoire de Neurobiologie, France 

TEDESCHI, BRUCE, Louisiana State University 

TELFER, JANICE, Wake Forest University 

THIBAULT, LAWRENCE, University of Pennsylvania 

TOTH, JOSEPH, Hunter College 

TRICAS, TIMOTHY, Washington University School of Medicine 

TWERSKY, LAURA, Hunter College 

TYTELL, MICHAEL, Wake Forest University 

UENO, HIROSHI, Rockefeller University 

UGORETZ, JOHN, La Jolla High School 

VERSELIS, VYTALITAS, Albert Einstein College of Medicine 



76 MARINE BIOLOGICAL LABORATORY 

WALTON, ALAN J., Oxford University, UK 

WANG, XIN-SHANG, Vassar College 

WEBB, CHRISTINA, University of California, Los Angeles 

WESTENDORF, Jcv NNE M., Duke University 

WHITTAKER, Josi Howard University 

WHITTEM; SE, University of Pennsylvania 

WILLI A ' :OME, Hunter College 

Wool I .-; C., California State University, Los Angeles 

ZAKEV ;, JANE, University of Illinois 

z, JOSEPH, Albert Einstein College of Medicine 
ZEA. SVEN E., University of Texas 

ZECEVIC, DEJAN, Institute of Biological Research, Yugoslavia 
ZHAO, ZHAE-YIONG, Baylor College 



YEAR-ROUND PROGRAMS 

BOSTON UNIVERSITY MARINE PROGRAM (BUMP) 

Director 
STRICKLER, J. RUDI 

Faculty (of Boston University unless otherwise indicated) 

ATEMA, JELLE TAMM, SIDNEY L. 

FREADMAN, MARVIN TAMM, SIGNHILD 

HUMES, ARTHUR G. (Emeritus) TIERNEY, ANN JANE 

SUMAN, DANIEL VALIELA, IVAN 

Staff (of Boston University unless otherwise indicated) 

CROMARTY, STUART SUNLEY, DANIEL 

DZIERZEWSKI, MICHELLE TAYLOR, MARGERY 

HAHN, DOROTHY VAN ETTEN, RICHARD 

LOHMANN, DENAH WOODWARD, HELEN 

Graduate students 

ALBER, MERRYL COULTER, DOUGLAS 

BANTA, GARY COWAN, DIANE 

BARSHAW, DIANA ELLIS, SARAH 

BORRONI, PAOLA ELSKUS, ADRIA 

CHEN, CHONG GALLAGER, SCOTT 

CORROTO, FRANK CLICK, STEPHEN 

COSTA, JOSEPH HANDRICH, LINDA 

Undergraduates 

BRAN, TERRENCE MULSOW, SANDOR 

BROWN, SIDNEY SCOTT, MARSHA 

CARLON, DAVID TAMSE, ARMANDO 

MURPHY, TARA TROTT, THOMAS 

HAHN, JILL WEBB, JACQUELINE 

HERSH, DOUGLAS WHITE, DAVID 

KRIEGER, YUTTA WOODS, SUSAN 

LAVALLI, KARI SAPONARO, STEPHEN 

MERCURIC, KIM SHAPIRO, RACHEL 

MERRILL, CARL WALLACE, RICHARD 
MOORE, PAUL 



RESEARCH AND TRAINING PROGRAMS 77 



Visiting investigators 

D'AVANSO, CHARLENE, Hampshire College 
POOLE, ALAN, Boston University 
RIETSMA, CAROL, SUNY, New Paltz 
SARDA, RAFAEL, University of Barcelona 
VOIGT, RAINER, University of Gottingen 



DEVELOPMENTAL AND REPRODUCTIVE BIOLOGY LABORATORY 
Director 
GROSS, PAUL R. 

LABORATORY OF BIOPHYSICS 
Director 
ADELMAN, WILLIAM J., JR. 

Staff (of NINCDS/NIH unless otherwise indicated) 
Section on Neural Membranes 

CLAY, JOHN R. 

FOHLMEISTER, JuRGEN R., University of Minnesota 

GOLDMAN, DAVID E., SUNY, Binghamton 

HODGE, ALAN J., Marine Biological Laboratory 

KRAMER, JUDITH A., University of Cincinnati College of Medicine 

LAVOIE, ROBERT, Marine Biological Laboratory 

MARTIN, DOROTHY L. 

McMAHON, WILLIAM E., Marine Biological Laboratory 

MUELLER, RUTHANNE, Marine Biological Laboratory 

RICE, ROBERT V., Carnegie Mellon University 

STANLEY, ELIS F. 

TYNDALE, CLYDE L., Marine Biological Laboratory 

WALTZ, RICHARD B., Marine Biological Laboratory 

Section on Neural Systems 

ALKON, DANIEL L., Chief 

BANK, BARRY, University of Toronto 

CHEN, CHONG 

COLLIN, CARLOS 

COULTER, DOUGLAS, Boston University 

DISTERHOFT, JOHN, Northwestern University Medical School 

HARRIGAN, JUNE, Marine Biological Laboratory 

HOPP, HANS-PETER 

IKENO, HIDETOSHI 

KUBOTA, MlCHINORI 

KUZIRIAN, ALANM. 

KUZIRIAN, JEANNE 

LEDERHENDLER, IZJA, Marine Biological Laboratory 

LEIGHTON, STEPHEN, Biomedical Engineering and Instrumentation Branch, NIH 

LOTURCO, JOSEPH 

McPHiE, DONNA 

NAITO, SHIGETAKA 

NEARY, JOSEPH, Marine Biological Laboratory 

SAKAKIBARA, MANABU 



78 MARINE BIOLOGICAL LABORATORY 

LABORATORY OF CARL J. BERG, JR. 
Director 
BERG, CARL J, JR. 

Staff 

ADAMS, NANCY 
ORR, KATKERINE S. 

Visiting investigators 

FARMER, MARY, Sea Education Association 

WARD, JACK, Division of Fisheries, Government of Bermuda 

LABORATORY OF CAROL L. REINISCH 

Director 

REINISCH, CAROL L., Tufts University School of Veterinary Medicine 

Staff 

MIOSKY, DONNA 
SMOLOWITZ, ROXANNA 

LABORATORY OF D. EUGENE COPELAND 
Director 
COPELAND, D. EUGENE 

LABORATORY OF DEVELOPMENTAL GENETICS 

Director 

WHITTAK.ER, J. RICHARD 
Staff 

CROWTHER, ROBERT 
LOESCHER, JANE L. 
MEEDEL, THOMAS H. 
MERCURIO, KIMBERLY 

Visiting investigators 
COLLIER, J. R., Brooklyn College 
Summer intern (undergraduate) 
ZELLER, ROBERT, Boston University 

LABORATORY OF JUDITH P. GRASSLE 
Director 

GRASSLE, JUDITH P. 
Staff 

GELFMAN, CECILIA E. 
MILLS, SUSAN W. 



RESEARCH AND TRAINING PROGRAMS 79 

LABORATORY FOR MARINE ANIMAL HEALTH 
Director 

LEIBOVITZ, Louis, Cornell University 
Staff 

ABT, DONALD A., University of Pennsylvania 
HAMILTON, HEATHER A., Cornell University 
JENNER, JENNIFER L., Cornell University 
McCAFFERTY, MICHELLE, Cornell University 
MONIZ, PRISCILLAC., Marine Biological Laboratory 

LABORATORY OF OSAMU SHIMOMURA 
Director 

SHIMOMURA, OSAMU, Boston University School of Medicine 
Staff 

SHIMOMURA, AKEMI 
Visiting investigators 

MUSICKI, BRANISLAV, Harvard University 
NAKAMURA, HIDESHL Harvard University 

LABORATORY OF RAYMOND E. STEPHENS 

Director 

STEPHENS, RAYMOND E., Marine Biological Laboratory/Boston University School of 
Medicine 

Staff 

GOOD, MICHAEL J., Marine Biological Laboratory 

OLESZKO-SZUTZ, SUSAN, Marine Biological Laboratory 

STOMMEL, ELIJAH W., Marine Biological Laboratory/Boston University School of Medicine 

LABORATORY OF SENSORY PHYSIOLOGY 
Director 
FEIN, ALAN 
Staff 

HAROSI, FERENC I. 
PAYNE, RICHARD 
SZUTS, ETE Z. 
WOOD, SUSAN 
ZAHAJSZKY, TIBOR 

Visiting investigators 

CORNWALL, CARTER, Boston University School of Medicine 
HAWRYSHYN, CRAIG W., Cornell University 
PETRY, HEYWOOD M., SUNY, Stonybrook 



80 MARINE BIOLOGICAL LABORATORY 

TSACOPOULOS, MARCO, University of Geneva, Switzerland 
WALZ, BERND, University of Ulm, West Germany 



LABORATORY OF SHINYA INOUE 

Director 

INOUE, SHFNYA, Marine Biological Laboratory, University of Pennsylvania 

Staff 

ANNIBALLI, DYON, Cornell Engineering School 
BOYD, STEVEN, Cornell Engineering School 
GREEN, DANIEL, Cornell Engineering School 
INOUE, THEODORE, Cornell Engineering School 
RUBINOW, JERRY, Cornell Engineering School 
SHIMOMURA, SACHI 
WOODWARD, BERTHA M. 



LABORATORY OF NEUROBIOLOGY 

Director 

REESE, THOMAS S. 

Staff (of NINCDS/NIH unless otherwise indicated) 

ANDREWS, S. BRIAN 

BURGER, TINA, Marine Biological Laboratory 

CHENG, TONI 

CHLUDZINSK.I, JOHN, Marine Biological Laboratory 

CRISE, BRUCE, Marine Biological Laboratory 

EVENDEN, PHYLLIS 

FROKJAER-JENSEN, JORGEN, University of Copenhagen 

GALLANT, PAUL 

GARBUS-GOOCH, CYNTHIA, Marine Biological Laboratory 

HAMMAR, KATHERINE 

JAROCHE, DEANNA, Marine Biological Laboratory 

KHAN, SHAHID, Marine Biological Laboratory 

MCCUSKER, ELIZABETH 

MURPHY, JOHN C., Marine Biological Laboratory 

REESE, BARBARA F. 

SHEETZ, MICHAEL P., Washington University 

SCHNAPP, BRUCE J. 

TATSUOKA, HOZUMI 

TERASAKJ, MARK 

VALE, RONALD D. 

WALROND, JOHN P. 

WISGIRDA, MARY, Marine Biological Laboratory 



NATIONAL FOUNDATION FOR CANCER RESEARCH 

Director 
SZENT-GYORGYI, ALBERT 



RESEARCH AND TRAINING PROGRAMS 8 1 

Staff 

GASCOYNE, PETER R. C. 

MCLAUGHLIN, JANE A. 

MEANY, RICHARD A. 

PETHIG, RONALD, University College of North Wales, UK 

Student 

PRICE, JONATHAN A., University College of North Wales, UK 

NATIONAL VIBRATING PROBE FACILITY 
Director 
JAFFE, LIONEL, Marine Biological Laboratory 

Staff 

DIXON, STEVEN 

SHIPLEY, ALAN 
STEWART, MARY 
WILLIAMS, PHILLIP C. 

Visiting investigators 

ALLEN, NINA, Wake Forest University 

BJORKMAN, THOMAS, Cornell University 

BOWDAN, ELIZABETH, University of Massachusetts, Amherst 

DURHAM, JOHN, Mt. Sinai Hospital, New York 

ETTENSOHN, CHARLES, Duke University 

FINK, RACHEL, Mount Holyoke College 

FLUCK, RICHARD, Franklin & Marshall College 

KATZ, URI, Israel Institute of Technology, Haifa, Israel 

KUNKEL, JOSEPH, University of Massachusetts 

LEVY, SIMON, Boston University 

PAYNE, RICHARD, Marine Biological Laboratory 

RUBIN, CLINTON, Tufts Medical School 

SARDET, CHRISTIAN, Station Marine Villelfranche sur Mer, France 

SKADHAUGE, ERIC, Royal Veterinary University, Copenhagen 

SPEKSNEIJDER, J. H., Marine Biological Laboratory 

TRINKAUS, JOHN, Yale University 

TROXELL, CYNTHIA, University of Colorado, Boulder 

WEIJER, KEES, University of Munich, FRG 

WEISENSEEL, MANFRED, University of Karlsruhe, FRG 

ZIVKOVIC, DANA, University of Utrecht 

THE ECOSYSTEMS CENTER 
Director 
HOBBIE, JOHN E. 

Staff and consultants 

BANTA, GARY GIBLIN, ANNE 

BOWLES, FRANCIS P. GRIFFIN, ELISABETH A. 

FERRY, ELIZABETH HELFRICH, JOHN V. K. 

GARRITT, ROBERT HOUGHTON, RICHARD A. 



82 MARINE BIOLOGICAL LABORATORY 

JOHNSON, STEPHEN POVIA, SANDRA 

LAUNDRE, JAMES RAY, ANDREA 

LEFKOWITZ, DANIEL REGAN, KATHLEEN 

MATHERLY, WALTHZR SEMINO, SUZANNE 

MCKERROW, ALFXA SHAVER, GAIUS R. 

MELILLO, JERRY M. STEUDLER, PAUL A. 

MICHENER, ROBERT STONE, THOMAS A. 

NADELHOFFER, KNUTE J. TUCKER, JANE 

OPPENHEIMER, JILL TURNER, ANDREA R. 

PETERSON, BRUCE J. WHITE, DAVID 

PLUMMER, NANCY YANDOW, TIMOTHY 

Trainees 

RASTETTER, EDWARD, University of Virginia 
RUDNICK, DAVID, University of Rhode Island 

Visiting scientists 

JORDAN, MARILYN J. 
O'BRIEN, W. JOHN 
RUBLES, PARKE 
WARING, RICHARD 

XI. HONORS 
FRIDAY EVENING LECTURES 

SIMBERLOFF, DANIEL, Florida State University, 27 June, "Academic Ecology and Environ- 
mental Problems: Red Scales, Vampire Bats, and Spotted Owls" 

ALBERSHEIM, PETER, University of Georgia, 4 July, "Oligosaccharins A New Class of Regu- 
latory Molecules in Plants and Animals" 

BROWN, DONALD D., Carnegie Institution of Washington, 1 1 July, "The Molecular Basis of 
Differential Gene Expression" 

STEVENS, CHARLES F., Yale University School of Medicine, 17, 1 8 July, Forbes Lectures, "Mo- 
lecular Basis for the Brain's Electrical Activity: I. Electrical Excitability of Neurons: 
II. Communication between Neurons" 

KAISER, DALE, Stanford University School of Medicine, 25 July, "Cell-Cell Interactions in a 
Simple Developmental Pathway" 

REESE, THOMAS S., NINCDS, NIH, and Marine Biological Laboratory, 1 August, "Kinesin 
An MBL Project" 

DOOLITTLE, RUSSELL F., University of California, San Diego, 8 August, "Evolution of the 
Vertebrate Plasma Proteins " 

KANDEL, ERIC R., College of Physicians & Surgeons of Columbia University and the Howard 
Hughes Medical Institute, 1 5 August, Lang Lecture, "The Long and Short of Memory" 

SELA, MICHAEL, The Weizmann Institute of Science, 22 August, "From Synthetic Antigens to 
Synthetic Vaccines" 

TRINKAUS, J. P., Yale University, 29 August, "Metazoan Cell Movements: Invasion and Mor- 
phogenesis" 

CHARLES ULRICK AND JOSEPHINE W. BAY FOUNDATION FELLOWSHIP 
SMOLOWITZ, ROXANNA, Marine Biological Laboratory 

ERNEST EVERETT JUST FELLOWSHIPS IN BIOLOGY 
JOSIAH MACY, JR., FOUNDATION 

WHITE, ROY L., Albert Einstein College of Medicine 



HONORS 83 

MBL SUMMER FELLOWSHIPS 

DLIBE, FRANCOIS, Universite du Quebec a Rimouski, Canada 

EHRLICH, BARBARA, University of Connecticut Health Center 

PIERSON, BEVERLY K., University of Puget Sound 

POOLE, THOMAS L., SUNY, Syracuse 

ROME, LAWRENCE C., University of Tennessee 

TAKEDA, KENNETH, University Louis Pasteur, France 

TRAVIS, JEFFERY, Vassar College 

BIOLOGY CLUB OF NEW YORK 
KASMER, JOHN M., University of Vermont 

FATHER ARSENIUS BOYER SCHOLARSHIP 
KASMER, JOHN M., University of Vermont 

GARY N. CALKINS MEMORIAL SCHOLARSHIP 
DIOGENE, GEORGE F., University of Barcelona, Spain 

FRANCES S. CLAFF MEMORIAL SCHOLARSHIP 
FREY, IRIS J. F., Philipps-University Marburg, FRG 

EDWIN GRANT CONKLIN MEMORIAL SCHOLARSHIP 
C/HARA, ELLEN M., Villanova University 

LUCRETIA CROCKER SCHOLARSHIP 

FALK, KATHLEEN, University of Massachusetts 
HART, ROBERTA., University of California, Berkeley 
MORUCCI, CARLO, University of La Sapienza of Rome, Italy 
ZAPATA, FERNANDO A., University of Arizona 

FOUNDERS-OTTO LOEWI 
AKINS, ROBERT E., JR., University of Pennsylvania 

FOUNDERS- WALTER E. GARREY 
C/HARA, ELLEN M., Villanova University 

FOUNDERS-S. O. MAST 
SMOLICH, BEVERLY, University of Virginia 

ALINE D. GROSS SCHOLARSHIP 
RENDER, JoANN, Hamilton College 

MERKEL H. JACOBS SCHOLARSHIP 
KASMER, JOHN M., University of Vermont 



84 MARINE BIOLOGICAL LABORATORY 

ARTHUR KLORFEIN FUND SCHOLARSHIPS 

HAMMER, MARTIN, Institut fuer Tierphysiologie, FRG 
HARRINGTON, MARY E,, Dalhousie University, Canada 
JURNISCH, VK ; . A.. University of California, Irvine 
LUSTIG, CORN'. I miann Institute, Israel 
SUPATTAPONF, MAI, Johns Hopkins University 

LUCILLE P. MARKEY CHARITABLE TRUST SCHOLARSHIPS 

BISWAS, SURAJIT, University of Pennsylvania 

BLOOM, THEODORA L., University of Cambridge, England 

BRADLEY, DAVID, University of Pennsylvania 

BROWN, ANNEC, University of Oregon 

CAULEY, KEITH A., University of Michigan 

DAHL, STEPHEN C., Wesleyan University 

DASSO, MARY C., Cambridge University, UK 

DEYST, KATHERINE A., Tufts University 

DIOGENE, GEORGE F., University of Barcelona, Spain 

DUBILIER, NICOLE, University of Hamburg, FRG 

FALK, KATHLEEN, University of Massachusetts 

FOLTZ, KATHLEEN R., Purdue University 

FREY, IRIS J. F., Philipps-University Marburg, FRG 

GANNON, PAMELA M., Tufts University 

GUDEMAN, DAVID M., Kansas University 

HAFNER, MATHIAS, German Cancer Research Center, FRG 

HART, ROBERTA., University of California 

HOULISTON, EVELYN, University of Cambridge, UK 

KOENIG, GERD, MPI fur Entwicklungsbiologie, FRG 

KUBIAK, JACEK Z., Warsaw University, Poland 

SAAVEDRA, CAROL, McGill University, Canada 

SMOLICH, BEVERLY, University of Virginia 

SVENDSEN, BETTY-ANN E., University of Dallas 

SYMES, KAREN, National Institute of Medical Research, UK 

TALEVI, RICCARDO, University of Naples, Italy 

THIVAKARAN, ALAGIRI G., Annamalai University 

VELLECA, MARK A., Yale University 

VITES, ANA M., University of Connecticut Health Center 

ZAPATA, FERNANDO A., University of Arizona 

ALLEN M. MEMHARD SCHOLARSHIP 
BROWN, ANNEC., University of Oregon 

JAMES S. MOUNTAIN MEMORIAL FUND, INC. SCHOLARSHIPS 1986 

DAHL, STEPHEN, Wesleyan University 

DASSO, MARYC., Cambridge University, UK 

FOLTZ, KATHLEEN R., Purdue University 

GUDEMAN, DAVID M., Kansas University Medical Center 

KATZ, KENNETH S., University of Massachusetts 

SYMONS, MARC H. C., Weizmann Institute, Israel 



HONORS 



85 



JAMESS. MOUNTAIN MEMORIAL FUND, INC. SCHOLARSHIPS 1985* 

CHEN, TUNG-LING, University of Maryland 
GOODWIN, ELIZABETH B., Brandeis University 
HANNA, MAYA, Harvard University 
PRET, ANNE-MARIE, Wesleyan University 
WALTHER, ZENTA, Yale University 
Wu, BEI-YUE, Wayne State University 

SOCIETY OF GENERAL PHYSIOLOGISTS SCHOLARSHIPS 

BLOOM, THEODORA L., University of Cambridge, UK 
HOULISTON, EVELYN, University of Cambridge, UK 

SURDNA FOUNDATION SCHOLARSHIPS 

SPANO, ANNAMARIA, Institute Superiore di Sanitz, Rome, Italy 
SVENDSEN, BETTY-ANN E., University of Dallas 

MARJORIE W. STETTEN SCHOLARSHIP 
SCHWEIZER, FELIX E., Biozentrum/Universitat Basel, Switzerland 



XII. INSTITUTIONS REPRESENTED 



U.S.A. 



Academy of Natural Sciences of 

Philadelphia 
Albany Medical Center 
Albert Einstein College of Medicine 
American Health Foundation 
Amherst College 
Arizona Research Laboratory 
Arizona State University 
Arizona, University of 
Arizona, University of, College of Medicine 
Atlantex and Zieler Instrument 

Corporation 
Axon Instruments, Inc. 
Bardeen Laboratory 
Bausch & Lomb 
Baylor College 
Baylor College of Medicine 
Beckman Instruments, Inc. 
Bell Laboratories 
Bethesda Research Labs 
Bigelow Laboratories 
BioTechnical International Inc. 
Biodyne Electronics 
Biomedical Engineering and 

Instrumentation Branch, NIH 
Bodega Marine Station 



Boston University 

Boston University Marine Program 

Boston University Medical School 

Bowling Green State University 

Bowman Gray Medical School 

Braintree Laboratories 

Brandeis University 

Brinkmann Instruments 

Brooklyn College 

Brown University 

California Institute of Technology 

California State University 

California State University, Los Angeles 

California, University of 

California, University of, Berkeley 

California, University of, Davis 

California, University of, Irvine 

California, University of, Los Angeles 

California, University of, Riverside 

California, University of, San Diego 

California, University of, San Francisco 

Carnegie Institution of Washington 

Carnegie-Mellon University 

Case Western Reserve University 

Center for Advanced Research 

Center for Astrophysics 



* The Marine Biological Laboratory regrets the omission of 1985 scholarship recipients in the 1985 An- 
nual Report [Biol. Bull. 171(1)]. 



86 



MARINE BIOLOGICAL LABORATORY 



Chesapeake Biological Laboratory 

Chicago, University of 

Chicago, University of, Medical School 

Childrens Hospital 

Cincinnati, University of, College of 

Medicine 

College of the Holy Cross 
Colorado, U mversity of 
Colorado, University of, Boulder 
Colorado Video 
Columbia University 
Columbia University College of Physicians 

and Surgeons 

Connecticut, University of 
Connecticut, University of. Health Center 
Connecticut, University of. Medical Center 
Connecticut, University of. School of 

Medicine 

Conservation Law Foundation 
Cornell Engineering School 
Cornell University 
Coulter Electronics 
Creighton University 
Crimson Camera Technical Sales, Inc. 
Dagan Corporation 
DAGE-MTI 
Dalhousie University 
Dallas, University of 
Damon Biotech, Inc. 
Dana-Farber Cancer Institute 
Dartmouth College 
Dartmouth Medical School 
Delaware, University of 
Denver, University of 
Dow Chemical 
Duke University 
Duke University Medical Center 
Dupont Corporation 
Earlham College 
Eastern Illinois University 
Eastman Kodak Company 
Emory University 

Emory University School of Medicine 
Environmental Protection Agency 
Ethicon, Inc. 
Evansville, University of 
Fairleigh Dickinson University 
Florida State University 
Florida, University of 
Flow Laboratory 
Fordham University 
Franklin and Marshall College 
General Electric Company 
General Scanning 

Georgetown University Medical School 
George Washington School of Medicine 



Georgia, University of 
Gilson Medical Electronics 
Gonzaga University 
Goucher College 
Grass Instrument Company 
Hacker Instruments 
Hampshire College 
Hahnemann University 
Hahnemann University School of 

Medicine 

G. W. Hannaway Associates 
Hartwick College 
Harvard Medical School 
Harvard University 
Harvard University School of Public 

Health 

Hawaii, University of 
Hinds Jr. College 
Hoefer Science Instruments 
Hope College 

Howard Hughes Medical Institute 
Howard University 
Hunter College 
Hutchinson Cancer Center 
IBI 
IDEA 

I.N.S.E.R.M. 
Ikegami Electronics Inc. 
ISCO 

Illinois Institute of Technology 
Illinois, University of, Chicago 
Illinois, University of. College of Medicine 
Illinois, University of, Urbana-Champaign 
Indiana University 

Instrumentation Marketing Corporation 
Interactive Video Systems 
International Business Machines 
Iowa, University of 
Johns Hopkins School of Medicine 
Johns Hopkins University 
Kansas City Veterans Administration 

Medical Center 
Kansas, University of 
Kansas, University of, Medical School 
Kenyon College 
Kip & Zonen 

Kisten, Babitsky, Latimer & Beitman 
Kresge Eye Institute 
LKB Instruments, Inc. 
Lab Line Instruments, Inc. 
LaFayette College 

Lamont-Doherty Geological Observatory 
Lander College, South Carolina 
Lehman College 
Leitz, E. Inc. 
Levity Corporation 



INSTITUTIONS REPRESENTED 



87 



Liberty Mutual Research Center 
Louisiana State University 
Louisville, University of 
META Systems, Inc. 
Maine, University of 
Mary Flagler Gary Arboretum, NY 
Maryland, University of 
Maryland, University of, Baltimore 
Massachusetts General Hospital 
Massachusetts Institute of Technology 
Massachusetts, University of 
Massachusetts, University of, Amherst 
Massachusetts, University of. Medical 

School 

Mayo Foundation 
Medical College of Ohio 
Medical College of Pennsylvania 
Memorial Sloan Kettering 
Merck, Sharp and Dohme Research 

Laboratories 
Miami, University of 
Miami, University of. School of Medicine 
Michigan State University 
Michigan, University of 
Millhauser Laboratory 
Minnesota, University of 
Miriam Hospital 
Missouri, University of 
Monsanto Company 
Mount Holyoke College 
Mount Sinai Hospital 
Mystic Marinelife Aquarium 
National Institute of Child Health and 

Human Development 
National Institute of Environmental Health 

Sciences 

National Institute of Mental Health/NIH 
National Institutes of Health 
National Institute of Neurological and 

Communicative Disorders and Stroke/ 

NIH 

National Marine Fisheries Service 
New Alchemy Institute 
New Brunswick Scientific, Inc. 
New England Medical Center 
New Hampshire, University of 
New Jersey Medical School 
New Jersey, University of. Medicine and 

Dentistry 

New York Blood Center 
New York, City University of 
New York Institute for Basic Research in 

Developmental Disabilities 
New York Institute for Basic Research in 

Mental Retardation 
New York Medical College 



New York, State University of, 

Binghamton 

New York, State University of, Buffalo 
New York, State University of, Downstate 

Medical Center 
New York, State University of. Health 

Sciences Center 

New York, State University of. New Paltz 
New York, State University of. Stony 

Brook 

New York University 
New York University College of Dentistry 
New York University Medical Center 
New York University School of Medicine 
Nichols College 
Nikon, Inc. 

North Carolina, University of 
North Nassau Mental Health Center 
Northeastern University 
Northwestern University 
Northwestern University Medical School 
Notre Dame, University of 
Oak Ridge National Laboratory 
Oberlin College 
Ocean Pond Corporation 
Oklahoma, University of 
Olympus Corporation of America 
Optiquip 
OPTRA, Inc. 
Oregon State University 
Oregon, University of 
Pennsylvania State University 
Pennsylvania, University of 
Pennsylvania, University of. School of 

Dental Medicine 
Pennsylvania, University of. School of 

Medicine 
Pharmacia, Inc. 
Photonic Microscopy 
Pittsburg, University of 
Portland State University 
Princeton University 
Procter and Gamble Company 
Puerto Rico, University of 
Puget Sound, University of 
Purdue University 
Quantex Corporation 
R & M Biometrics, Corp. 
Radiomatic Instruments 
Rainbow Babies and Children's Hospital 
Rainin Instrument Company 
Reed College 

Rensselaer Polytechnic Institute 
Rhode Island College 
Rhode Island, University of 
Rice University 



MARINE BIOLOGICAL LABORATORY 



Rochester, University of 

Rochester, University of. Medical Center 

Rochester, University of, School of 

Medicine and Dentistry 
Rockefeller Foundation 
Rockefeller University 
Rush Mecli .-;.;! Center 
Rush-pvc' : :>tcnan, St. Luke's Medical 

Center 

Rutgers University 
Rutgers University Medical School 
Savant Instruments 
Sea Education Association 
Simmons College 
Smith College 
Smithsonian Institution 
Sorvall Instruments 
South Carolina, University of 
Southeastern Massachusetts University 
Southern California, University of 
Stanford University 
St. Peter's College 
Swift Instruments 
Syntex 

Syracuse University 
Technical Products International, Inc. 
Temple University 
Temple University Medical School 
Tennessee, University of 
Tennessee, University of, Oak Ridge 
Texas Christian University 
Texas, University of 
Texas, University of, Austin 
Texas, University of. Health Center 
Texas, University of. Medical Branch 
Texas, University of. Medical School 
Texas, University of, Medicine and 

Dentistry 

Thomas Jefferson University 
Trinity College 
Tufts University 
Tufts University, Sackler School 
Tufts University School of Medicine 
Tufts University, School of Veterinary 

Medicine 



Union College 

United States Food and Drug 

Administration 
Upjohn Company 
Universal Imaging Corporation 
Upstate Medical Center 
VWR Scientific 
Vanderbilt University 
Vassar College 
Vermont, University of 
Veterans Administration Hospital, San 

Francisco 

Veterans Administration Medical Center 
Villanova University 
Virginia, University of 
B. Vittor and Associates 
Wake Forest University 
Washington and Lee University 
Washington State University 
Washington University 
Washington, University of 
Washington University School of Medicine 
Wayne State University 
Wesleyan University 
Whitman College 
Whitney Marine Laboratory 
Wilkes College 

William and Mary, College of 
Williams College 
Wisconsin, University of 
Wisconsin, University of, Madison 
Wistar Institute 
Woods Hole Data Base 
Woods Hole Education Associates 
Woods Hole Oceanographic Institution 
Woods Hole Research Center 
Worcester Foundation for Experimental 

Biology 

World Precision Instruments 
Yale University 

Yale University Medical School 
Carl Zeiss, Inc. 



FOREIGN INSTITUTIONS 



Alberta, University of, Canada 
Annamala University, India 
Barcelona, University of, Spain 
Basel, University of, Switzerland 
Bedford Institute of Oceanography, Canada 
Bergen, University of, Norway 
Calgary, University of, Canada 
Cambridge University, UK 



Centre National de la Recherche 

Scientifique, France 
Centra de Investigacion y de Estudios 

Avanzados del IPN, Mexico 
Copenhagen, University of, Denmark 
Dalhousie University, Canada 
Division of Fisheries, Bermuda 
Edinburgh, University of, Scotland, UK 



INSTITUTIONS REPRESENTED 



89 



Free University of Berlin, FRG 
Gadjah Mada University, Indonesia 
Geneva, University of, Switzerland 
German Cancer Research Center, FRG 
Hamburg, University of, FRG 
Hebrew University, Israel 
Heidelburg, University of, FRG 
Ibadan, University of, Nigeria 
I.D.E.A., Venezuela 
Institute of Animal Physiology, FRG 
Institute of Biological Research, Yugoslavia 
Institute de Investigacion Medica, 

Argentina 
Institute Superiore de Sanita of Rome, 

Italy 
Institute Venezolanode Investigaciones 

Cientifican, Venezuela 
International Laboratory for Research on 

Animal Diseases, Kenya 
Israel Institute of Technology, Israel 
Karlsruhe, University of, FRG 
Konstanz, University of, FRG 
La Sapienza of Rome, University of, Italy 
Laboratoire de Neurobiologie, France 
Leeds, University of, UK 
Leuven, University of, Belgium 
Liverpool, University of, UK 
Lund, University of, Sweden 
Marie Curie, University of, France 
Max Planck Institut fur Cell Biologic, 

Heidelberg, FRG 
McGill University, Canada 
McMaster University, Canada 
Milan, University of, Italy 



Munich, University of, FRG 

Naples, University of, Italy 

National Institute of Basic Biology, Japan 

National Institute of Medical Research, 

UK 

Osaka University, Japan 
Ottawa, University of, Canada 
Oxford University, UK 
Panum Institute, Denmark 
Philipps-University, Marburg, France 
Quebec, University of 
Queen's University, Canada 
Royal Veterinary University, Denmark 
Ruhr-Universitat Bochum, FRG 
Seville, University of. Medical School, 

Spain 

Statens Serum Institute, Denmark 
Station Marine Villefranch sur Mer, France 
Station Zooligique, France 
Stazione Zoologica, Naples, Italy 
Stockholm, University of, Sweden 
Toronto, University of, Canada 
Tottori University, Japan 
Universidad de Cuyo-Conicet, Argentina 
Universite Louis Pasteur, France 
University College, London, England, UK 
University College, Northern Wales, UK 
University College, Oxford, UK 
Uppsala University, Sweden 
Walter and Eliza Hall Institute, Australia 
Warsaw University, Poland 
Weizmann Institute of Science, Israel 
Wellcome Laboratories, UK 



XIII. LABORATORY SUPPORT STAFF 

Including Persons Who Joined or Left The Staff During 1986 
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METZ, CHARLES B., Editor MOUNTFORD, REBECCA J. 

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90 



MARINE BIOLOGICAL LABORATORY 



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ENOS, EDWARD G., JR. 
ENOS, JOYCE B. 



LABORATORY SUPPORT STAFF 



91 



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FRANK, DONALDS. 
HANLEY, JANICE S. 

Public Information Office 

SHREEVE, JAMES M., Director 
LILES, GEORGE W., JR., Director 
CROSBY, CAROL 
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Research Services 

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MARTIN, LOWELL V. 

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Sponsored Programs 

HOWARD, JOAN E., Coordinator 
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DWANE, FLORENCE 

1986 Summer Support Staff 

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ASHMORE, LYNNE E. 
BELIVEAU, CHRISTINE A. 
BERG, CARL J., Ill 
BINDA, JOHN H. 
BURTON, RICHARD W. 
CHILDERS, REBECCA L. 
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DEMELLO, KIMBERLY A. 
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DONOVAN, JASON P. 
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TASSINARI, EUGENE 



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PAUK, CHRISTINE 
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to Electron Microscope Laboratory 
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POTHIER, JAHN A. 
PRINDLE, KIRK 
REMSEN, ANDREW W. 
RICHARDSON, KEITH W. 
ROSE, CHRISTINE 
ROZANSKI, CHRISTOPHER 
SANGER, RICHARD H., JR. 
SHEEHAN, PETER C. 
SINAGRA, DAVID T. 
SLOANE, MICHAEL B. 
SWOPE, JOHN G. 
VALOIS, FRANCIS X. 
VANALSTYNE, MARK 
WETZEL, ERNEST D. 
WHEELER, BRADLEY E. 
WINSPEAR, DAVID A. 
WYTTENBACH, ANN G. 



Reference: Biol. Bull. 173: 92-109. (August, 1987) 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS AND OTHER 
EICOSANOIDS IN INVERTEBRATES 

DAVID W. STANLEY-SAMUELSON 

Dcj'ii.'-nnent of Entomological Sciences, University of California, Berkeley, California 94720 

ABSTRACT 

Prostaglandins and other biologically active derivatives of polyunsaturated fatty 
acids have been detected in a large number of invertebrate species. A brief summary 
of the mammalian background of arachidonic acid metabolism is provided, and the 
physiological significance of these compounds in invertebrates is reviewed. Topics 
include regulation of ion flux, temperature regulation, reproductive biology, cell ag- 
gregation, and host-parasite interactions. Finally, perspectives on current and possi- 
ble future research are offered. 

INTRODUCTION 

The term eicosanoid was introduced and used by Corey et al. (1980) to describe 
the various biologically active derivatives of eicosapolyenoic fatty acids, especially 
arachidonic acid. So far, we know of four major groups of eicosanoids: the prostaglan- 
dins (PCs), the hydroperoxy- and hydroxyeicosatetraenoic acids (HPETEs and 
HETEs), the leukotrienes (LTs), and the lipoxins (LXs). Interest in the significance 
of eicosanoids in the biology of mammals stems from physiological studies conducted 
in the early twentieth century. In the earliest reference to one group of eicosanoids, 
the PGs, Jappelli and Scafa (1906) noted that extracts of dog prostrate glands caused 
paralysis of central respiratory control and changed heart rates when injected into 
dogs and rabbits. The discovery of PG pharmacological activity in human seminal 
fluids (Kurzrok and Lieb, 1930) probably marks the beginning of the detailed studies 
of the clinical significance of these compounds. Elucidation of the chemical structures 
of PGs in the early 1960's (Bergstrom et al., 1962a, b) greatly increased the pace of 
research and discovery, hindered in that decade mainly by the limited availability of 
working quantities of purified compounds. It is now known that PGs are present and 
play important roles in almost all mammalian tissues and fluids (Horrobin, 1978). 
Examples of PG action include pathophysiological actions such as mediation of the 
inflammatory response (which we commonly block by ingestion of aspirin) and par- 
ticipation in the blood-clotting cascade, as well as physiological actions such as con- 
traction of smooth muscle. 

The growth of PG research began with initial physiological observations, along 
with isolation and structural determinations of individual PGs. This was followed by 
the development of techniques to produce PGs in a commercially profitable way for 
clinical and biological studies. Commercial production of PGs evolved from biosyn- 
thesis from appropriate precursor fatty acids using large-scale enzyme preparations, 
through the discovery of naturally occurring sources of PGs and of intermediates in 
chemical synthesis to economical total synthesis. Hence, the first report of PGs in 

Received 15 April 1987; accepted 26 May 1987. 

Abbreviations: PG == prostaglandin, LT = leukotriene, HETE = hydroxyeicosatetraenoic acid, 
HPETE = hydroperoxyeicosatetraenoic acid, LX = lipoxin. 

92 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 93 

an invertebrate animal, the gorgonian coral Plexaura homomella (Weinheimer and 
Spraggins, 1969), met with tremendous interest, not as a zoological discovery, but as 
a commercial source of PG for laboratory study. In the years between this first discov- 
ery of a potentially economical source of PG and the development of appropriate 
synthetic strategies, the search for other biological sources of PGs turned up many 
examples of their occurrence in marine invertebrates, albeit at tissue concentrations 
far below the point of commercial interest. 

One of the PGs in greatest abundance in the coral. 1 5-epi-PGA 2 , is not pharmaco- 
logically active in the usual mammalian biological assays for PG activity (Nakano, 
1969). Chemical modification of the naturally occurring form to clinically useful 
structures, as well as commercial, ecological, and environmental aspects of sustained 
PG yield from coral have been reviewed (Theoder, 1977; Berte, 1981; Bundy, 1985). 
Other papers describe evidence for the occurrence of PGs in over one hundred inver- 
tebrate species. Christ and van Dorp (1972) detected PG-biosynthesis activity in five 
invertebrates including two coelenterates, a mollusc, an annelid and an arthro- 
pod but not in two insect species. Using a classical bioassay for the pharmacological 
effect of PG on contraction of mammalian smooth muscle, Nomura and Ogata 
(1976) detected PGs in a procordate, and in representatives of Echinodermata, Mol- 
lusca, Annelida, Coelenterata, and Arthropoda (including an insect). PGs were also 
detected by bioassay in saliva of another terrestrial arthropod, the tick Boophilus 
microplus (Dickinson et al., 1976; Higgs et ai, 1976). Using radioimmunoassay, 
Shemesh et al. (1979) found PGs in reproductive organs and salivary glands of an- 
other tick. Since all PGs are formed from a common intermediate, prostaglandin- 
endoperoxide, PG synthesis could be inferred from activity of prostaglandin-endo- 
peroxide synthetase. Morse et al. (1978) detected this enzyme activity in 41 species 
of coelenterates collected in the Caribbean Sea and the Pacific Ocean. Gromov et al. 
(1982) used radioimmunoassay to estimate amounts of two PGs in a snail. Korot- 
chenko et al. (1983) found smooth muscle-contracting activity in 10 echinoderm 
species; they also refer to finding PG activity in 40 other invertebrates. 

Aside from detection of PGs in a large number of invertebrate species, certain 
reports suggest that eicosanoids play fundamental physiological roles in representa- 
tives of many invertebrate phyla. Such findings are interesting because they provide 
insights into the details of regulatory physiology. Interest extends to an evolutionary 
axis because discovery of eicosanoid physiology especially in the very early phyla 
suggests that the significance of these compounds is not limited to vertebrate and 
clinical physiology, but was established early in metazoan evolution. 

Evolutionary interest may eventually extend to plants, as well. Gregson et al. 
(1979) described the occurrence of two PGs in the red alga Gracilaria lichenoides, 
and Janistyn (1982) reported chemical identification of PGF 2a in the flowering plant 
Kalanchoe blossfeldiana. A prostaglandin-like compound was produced from lin- 
olenic acid by an enzyme preparation of flaxseed (Zimmerman and Feng, 1978). The 
physiological significance of these compounds in plants is not clear, but compounds 
that inhibit PG-biosynthesis in mammals inhibited growth in four fungus species 
(Herman and Herman, 1985; Kerwin et al., 1986). Earlier inhibitor studies showed 
inhibition of flowering in Pharbitis nil (Groenewald and Visser, 1974). Although 
these findings are preliminary, they suggest that eicosanoids may be of broad biologi- 
cal significance. 

The goal of this review is to provide an appreciation of the physiological signifi- 
cance of eicosanoids in invertebrate animals. Since the appropriate nomenclature 
and physiological background comes from decades of work on various mammal sys- 
tems, it is useful to begin with a background from mammal studies. 



94 D. W. STANLEY-SAMUELSON 

A BACKGROUND FROM MAMMALIAN STUDIES 

Upon stimulation by various agonists, many mammal cells hydrolyze polyunsat- 
urated fatty acids (PUFAs), by action of phospholipase A2, from the beta carbon of 
membrane phospholipids. Three C20 PUFAs dihomo-7-linolenic (C20:3n6), ara- 
chidonic (C20:4n6), and eicosapentaenoic (C20:5n3) acids may be metabolized by 
one of two major pathways into biologically active molecules. In the cyclooxygenase 
pathway, PUFAs are transformed into prostaglandins and thromboxanes, whereas 
the lipoxygenase pathway leads to hydroperoxy- and hydroxypolyenoic fatty acids 
which are themselves biologically active as well as further metabolized into lipoxins 
and leukotrienes. Since these are all derivatives of C20 PUFAs, they may be collec- 
tively referred to as eicosanoids. The following description of the biosynthesis and 
physiological roles of these compounds in mammals is assembled from several re- 
views and books (Horrobin, 1978; Samuelsson el a/., 1978; Hansson el al, 1983; 
Samuelsson, 1983; Serhan el al., 1985), and is presented with minimum referencing. 

PGs are C20 carboxylic acids with a five-membered ring variously substituted at 
C9 and Cl 1, and two aliphatic chains featuring a substitution at C15 and one, two, 
or three double bonds. The structures of the principle PGs are shown in Figure 1 . PGs 
are designated as lettered and numbered series. The numbers indicate the number of 
aliphatic double bonds, giving rise to the one-, two-, and three-series PGs. The letters 
are associated with the particular pattern of substitutions on the five-membered ring: 
PGE features C9 keto, Cl 1 hydroxyl substitutions; PGF a C9,C1 1 dihydroxyl pat- 
tern; PGD a C9 hydroxyl, Cl 1 keto arrangement. PGs of the A, B, D, E, and F series 
are so distinguished. 

Biosynthesis of PGs is a multistep operation beginning with formation of the pros- 
taglandin endoperoxides first PGG by action of microsomal prostaglandin endo- 
peroxide synthetase. The same enzyme also cleaves the hydroperoxy group of PGG 
to form PGH. PGH is the root intermediate in the synthesis of the primary PGs: PGD 
is formed by a glutathione-S-transferase, PGE requires prostaglandin endoperoxide 
E isomerase and PGF prostaglandin endoperoxide reductase; PGI is formed by pros- 
taglandin endoperoxide I isomerase and thromboxane A (TxA) by prostaglandin 
endoperoxide thromboxane A isomerase. 

PGs have been detected in most mammalian tissue systems where they are in- 
volved in many well-catalogued (Horrobin, 1978) physiological activities. Examples 
of PG action include contraction of smooth muscle (i.e., uterine, gut, and blood ves- 
sel), attenuating cellular response to hormones, and release of digestive acid in the 
stomach. Thromboxane A 2 is a potent inducer of platelet aggregation; its name is 
taken from its origin, the thrombocytes. 

Lipoxygenase pathways first lead to hyproperoxy fatty acids which can be reduced 
by peroxidases, and possibly by non-enzymatic reactions, to corresponding hydroxy 
fatty acids (Fig. 2). Arachidonic acid is the best-studied lipoxygenase substrate in 
mammals, and oxygen can be added at various positions, leading to 5-, 8-, 9-, 1 1-, 
12-, and 1 5-hydroxyeicosatetraenoic acids (the various HETEs). Di- and tri-hydroxy 
fatty acids also can be formed by lipoxygenase acting on the same fatty acid substrate 
more than once; another route to trihydroxy acids is by way of an epoxy-hydroxy 
acid. While PGs are involved in various physiological as well as pathophysiological 
actions, the lipoxygenase products apparently are involved in pathophysiological ac- 
tions such as bronchial constriction. The lipoxygenase reactions are found in defense 
systems such as the various leukocytes, macrophages, monocytes, lung, and spleen. 
HETEs are biologically active in defense roles. For example, 5-, 9-, and 1 1-HETE are 
all active in inducing the chemokinesis and chemotaxis associated with migration of 
eosinophils into the site of certain hypersensitivity reactions. 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 95 



PROSTAGLANDINS 




homo-tf-LINOLENIC OH 



OH 




ARACHIDONIC 




EICOSAPENTAENOIC OH 



RING FEATURES OF PROSTAGLANDINS 




PGA 



FIGURE 1 . Relationship between the 1 -, 2-, and 3-series prostaglandins and their parental polyunsat- 
urated fatty acids, respectively C20:3n6, C20:4n6, and C20:5n3, is indicated by the arrows. X indicates 
cyclooxygenase activity. Ring features of five prostaglandins are shown in the lower panel where R stands 
for the aliphatic chains shown on the complete structures. 



The leukotrienes (LTs; Fig. 3) were discovered during work on rabbit polymor- 
phonuclear leukocytes, and take their names from this and the conjugated triene 
structure they have in common. The following description of LTs comes from the 
review by Samuelsson (1983). There are two classes of leukotrienes: the cysteine- 
containing group (LTC 4 , LTD 4 , and LTE 4 ), and LTB 4 , which is not substituted. 
Biosynthesis of the LTs begins with formation of 5-hydroperoxy-6,8,l 1,14-eicosa- 
tetraenoic acid (5-HPETE) by action of lipoxygenase followed by conversion to LTA 4 
by abstraction of a hydrogen and elimination of a hydroxyl anion, catalyzed by a 
soluble enzyme, dehydrase. LTA 4 is converted to LTB 4 by hydrolase, or into the 
parental cysteine-containing LT (LTC 4 ) by a glutathione-S-transferase. The cysteine- 
containing LTs feature a thioether linkage at C6 to cysteine; LTC 4 is 7-glutamyl- 
cysteinyl-glycyl substituted; glutamyl transpeptidase elimination of the glutamine 
residue forms cysteinylglycyl LTD 4 which can be metabolized into cysteinyl LTE 4 . 

LTs have been identified in several cell systems including rabbit, human, mouse, 
and rat leukocytes; mouse and rat macrophages; and human and guinea pig lung. 
The biological significance of these compounds lies in their identification as the slow- 
reacting substance of anaphylaxis (SRS-A). This material is a mediator in asthma and 
other mammalian hypersensitivity reactions; SRS-A is released with other mediators 



96 



D. W. STANLEY-SAMUELSON 



0(0)H 



COOH 




,COOH 



0(0)H 

15-H(P)ETE 



6-ri(P)ETE 



H(0)0 



H(0) 



.COOH 




COOH 



12-H(P)ETE 



,COOH 

9-H(P)ETE 0(0)H 

I1-H(P)ETE 

FIGURE 2. Structures of lipoxygenase metabolites of arachidonic acid, hydroxyeicosatetraenoic and 
hydroperoxyeicosatetraenoic acids. 



,COOH 



Arachidonic acid 



OOH 



LTA, 



,COOH 



5-HPETE 




Addition of, 
glutathione/ 



, Enzymatic 
^hydrolysis 



LTC 




A k= 



LTD, 




LTE, 




COOH 



CHCONHCH,COOH 
I i 

NHCOCH,CH 9 CHCOOH 

1 L I 

NH 



COOH 



CHCONHChUCOOH 

I i 

NH-, 



COOH 



CHCOOH 



.COOH 




LIB, 



FIGURE 3. Structures of leukotrienes. 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 97 

,COOH 

Arachidonic acid 





COOH 

15-HPETE 



HO OH H OH 

'COOH \ W rOOH 





H OH HO OH 

Lipoxin A (LX-A) Lipoxin B ( LX-Bl 

FIGURE 4. Structures of lipoxins. 



after interaction of antigens such as pollen with immunoglobulin. SRS-A is a mixture 
of the cysteine-containing LTs. LTB 4 , which does not contain cysteine, stimulates 
enzyme release, adhesion of neutrophils to endothelial cells, and movement of fluids 
through vessel walls in microcirculation. Lindgren el al. (1985) showed that LTs oc- 
cur in the rat brain most prominently in the hypothalamus and median emi- 
nence and that they may be involved in hormone release by brain cells. 

The lipoxins (Lx; Fig. 4) are the most recently discovered metabolites of arachi- 
donic acid. They share the characteristic feature of a conjugated tetraene structure. 
Two major LXs, LXA and LXB, were formed by human leukocytes; LXA stimulated 
oxygen metabolism and generation of active oxygen species in human neutrophils. 
The action of LXA in neutrophils differs from the action of leukotriene B 4 and may 
represent another physiological mechanism of host defense. Lipoxins appear to be 
formed by 5-lipoxygenase activity on a substrate formed by 1 5-lipoxygenase metabo- 
lism of arachidonic acid. (The trivial name lipoxins is an abbreviation of lipoxygenase 
interaction products.) 

The PGs, LTs, and LXs are involved in basic physiological processes at the cellu- 
lar level and appear to be especially important in various pathophysiological re- 
sponses such as inflammation, blood-clotting, asthma, and tumor growth. Due to 
their clinical significance, much effort is directed toward appreciating the regulation 
of arachidonic acid metabolism and developing specific inhibitors of PG, LT, and 
LX biosynthesis. Specific compounds will be mentioned in the contexts of biological 
studies in various invertebrate systems. Here it should be mentioned that within a 
given mammalian system there is considerable tissue variation in the effects of vari- 
ous inhibitors; moreover, there is variation between mammalian species. In light of 
tissue and specific variations in cyclooxygenase and lipoxygenase systems in mam- 
mals, one notes that the considerable literature on mammals should not be taken as 
a set of rules of the biochemistry of fatty acids in invertebrates. It is more appropriate 
to interpret the background as a loose set of guidelines, likely to be misleading at 



98 D. W. STANLEY-SAMUELSON 

crucial points in our consideration of the physiological significance of eicosanoids in 
invertebrates. 

PHYSIOLOGICAL SIGNIFICANCE OF EICOSANOIDS IN INVERTEBRATES 

Regulation of ion flux 

Like other freshwater bivalves, Ligumia subrostrata maintains its body fluids hy- 
perosmotic to the aquatic medium, largely by regulating the flux of sodium, its major 
blood cation (Dietz, 1977, 1979). PGE 2 appears to be a component of the sodium 
regulation system because inhibition of endogenous PG-biosynthesis by injection of 
indomethacin, a potent cyclooxygenase inhibitor in mammals, increased sodium 
flux. The effect lasted about 15 hours, approximately doubling the control values. 
Alternatively, when PGE 2 was injected in parallel experiments, sodium influx de- 
clined about 5-fold from control values. Since chloride concentrations and sodium 
outflux remained unchanged during these experiments, Graves and Dietz (1979) 
concluded that PGE 2 participates in ion regulation by specifically controlling so- 
dium influx. A tissue specificity may also exist because indomethacin modified the 
activity of the epithelial cells involved in sodium uptake without changing urinary 
sodium loss. 

Indomethacin stimulated sodium influx in a dose-dependent way over the con- 
centration range of 0.05 to 0.25 ^mol/g dry wt. Other PG-synthetase inhibitors in 
mammals meclofenamate (a cyclooxygenase inhibitor), and dexamethasone 
(which inhibits phospholipase A 2 , and hence, regulates substrate availability) also 
stimulate sodium uptake (Saintsing and Dietz, 1983). The stimulatory effect of PG- 
synthetase inhibitors was neutralized by co-injection of PGE 2 , supporting the view 
that PG is part of the system regulating epithelial sodium flux. PGE 2 reduces influx; 
reduction of PGE 2 biosynthesis may increase influx by attenuating the PG inhibition 
of uptake, but positive stimulation seems to depend on a biogenic amine, 5-hydroxy- 
tryptamine (5-HT, or serotonin), rather than on another PG since PGF 2a acts much 
like PGE 2 (Saintsing and Dietz, 1983). Cyclic AMP (cAMP) also stimulates sodium 
uptake (Graves and Dietz, 1982), which suggests that PG inhibition and 5-HT stimu- 
lation of sodium flux may both function via antagonistic effects on adenyl cyclase 
activity. 

Arachidonic acid injections apparently increased renal outflux of sodium without 
changing epithelial uptake. Graves and Dietz (1979) suggested that the arachidonic 
acid may initially alter renal function, and be metabolized too quickly to allow forma- 
tion of inhibitory levels of PGE 2 in epithelial tissue. Another possibility is that ion 
regulation is more complex (Graves and Dietz, 1982). If, as in mammals, arachidonic 
acid is potentially metabolized into a variety of prostanoid compounds, then we can 
imagine one metabolite, PGE 2 , inhibiting epithelial uptake while others, not yet iden- 
tified, modify renal ion flux in ways still unknown. 

The idea that PGs regulate epithelial sodium uptake in a freshwater mussel is 
based mostly on pharmacological treatments with appropriate compounds. Saintsing 
et al. (1983) showed the presence of PGs in L. subrostrata extracts by RIA, lending 
further support to natural occurrence and biological activity in an aquatic inver- 
tebrate. 

PGE 2 is also involved in ion regulation in the marine bivalve Modiolus demissus 
(Freas and Grollman, 1 980). When isolated gills were subjected to hypoosmotic stress 
by incubation for 60 minutes in 25% seawater, there was a 10-fold increase in PGE 2 
released into the medium, suggesting an increase in biosynthesis and release of the 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 99 

PG. In addition to this osmotic action on PG release, there is a specific ionic effect. 
To test for possible ionic effects, gills were incubated in artificial seawater of fixed 
osmotic concentration, but selectively free of sodium, calcium, potassium, or magne- 
sium. Only the magnesium-free artificial seawater stimulated gills to increase PGE 2 
release. However, the apparent osmotic effect is not due solely to depletion of envi- 
ronmental magnesium because gills incubated in hypoosmotic seawater with normal 
magnesium concentrations also induced increased PG release. Hence, gill tissues of 
this marine bivalve respond to changes in osmotic and ionic concentrations. 

In mammals, the physiological activities of many PGs are mediated by specific 
cellular receptor sites. Freas and Grollman (1981) showed the existence of specific 
PGA 2 binding sites in homogenates of gills, mantles, siphons, adductor, and upper 
and lower visceral masses. In gills, these sites were ionic, pH dependent, and revers- 
ible. To date this is the only study of PG binding sites in invertebrate tissues; such a 
finding adds considerable verisimilitude to physiological propeties of PGs. 

Mediation of behavioral thermoregulation and fever 

PGE, appears to mediate febrile response to infection in a number of mammals, 
including monkeys (Crawshaw and Still, 1975), sheep (Hales et ai, 1973), rabbits 
(Stitt, 1973; Lin, 1978), cats (Milton and Wendlandt, 1970; 1971), and guinea pigs 
(Szekely and Komaroni, 1978). Fever also occurs in non-mammalian vertebrates, 
although the increased body temperatures appear to be mediated by behavioral as 
opposed to endogeneous physiological mechanisms. Behavioral fever has been ob- 
served in frogs (Casterlin and Reynolds, 1977a, Myhre et ai, 1977), a lizard (Bern- 
heim and Kluger, 1976), and several fishes (Reynolds et al., 1976). 

Some aquatic invertebrates express behavioral fever in response to bacterial infec- 
tion by moving into a zone of warmer water. The freshwater crayfish Cambarus bar- 
toni exhibited a 2C behavioral fever after innoculation with a suspension of killed 
bacteria (Aeromonas hydrophila) by choosing higher temperatures in a gradient 
trough (Casterlin and Reynolds, 1977b). This behavioral response to infection may 
be mediated by endogenous formation of PGE, because increasing doses of the PG 
also induced 1 to 3.5C fevers when injected over the range of 50 to 500 ^/individual 
(Casterlin and Reynolds, 1978). Three marine arthropods the American lobster 
Homarus americanus, the pink shrimp Penaeus duorarum, and the horseshoe crab 
Limulus polyphemus similarly increased their temperature preferenda by more 
than 4C in response to 100 ^g injections of PGE, (Casterlin and Reynolds, 1979). 

Two terrestrial arthropods, the scorpions Bathus occitanus and Androctonnus 
australis, regulated their body temperatures by selecting appropriate positions along 
temperature gradients in a sand box. A. australis increased temperature preferences 
by 15C and B. occitanus by 20C after treatment with physiological doses of PGE, 
(Cabanac and Le Guelte, 1980). Although it is not known whether these species gen- 
erate fever due to bacterial infection, it appears that PGs may be involved in some 
aspect of behavioral thermoregulation. 

Together, these reports suggest that PGs may be some part of the thermoregula- 
tory physiology of many invertebrates. The idea is based on the observation of in- 
creased body temperatures in response to individual doses of a single compound, 
namely PGE,. Important detailed biochemical questions remain unanswered: do 
PGs naturally occur in these species? Does PG biosynthesis increase after infection, 
but before the febrile response? Do all PGs induce fever, or is a more specific set of 
these compounds involved? Research in this area may assume ecological interest, as 
suggested by remarks below. 



100 D. W. STANLEY-SAMUELSON 

Among terrestrial invertebrates, many medium to large size insects regulate tho- 
racic temperaiur <o a set point suited to the high metabolic demands of powered 
flight by belr ! (Casey, 1981) or physiological (Kammer, 1981) means. In addi- 
tion to flyin,: >, thermoregulation has been studied in caterpillers of two sphinx 
moths, // :/ and Manduca sexta (Casey, 1976, 1977). H. lineata appears to 
sustai x>dy temperatures and correspondingly high rates of feeding by basking 
in appro e postures; M. sexta does not maintain high temperatures even though 
feeding and growth rates are reduced considerably at cooler temperatures. These 
dirll-ivnt behaviors appear to be linked to differences in predator defense mechanisms 
and in seasonal availability of their host plants. Other caterpillers, including the but- 
terflies Vanessa io and V. urtica, huddle in groups, resulting in increased body tem- 
perature and development rates (Mosebach-Pukowski, 1938). Similarly, the larvae of 
wax moths thermoregulate, partly, by huddling or scattering (Smith, 1941). 

Many insect species are resistant to viral infection when maintained at higher 
temperatures (Tanada, 1967). Watanabe and Tanada (1972) reviewed several lepi- 
dopteran cases of insect viruses which do not cause lethal infections at higher temper- 
atures, including larvae of the armyworm Pseudaletia unipunctata, the cabbage 
looper Trichoplusia ni, and the corn ear worm Heliothis zea. Hence, behavioral ther- 
moregulation in invertebrates may effect such biological parameters as feeding and 
development rates, and resistance to disease. PGs may be an important biochemical 
mediator in this area of physiological ecology. 

Control of hatching 

In the barnacle Balanus balanoides, full egg-laying involves passing eggs along 
oviducts into ovisacs produced by oviducal glands. Fully formed egg masses are fi- 
nally released into the mantle cavity, where they remain until hatching which corre- 
sponds with spring algal blooms (Crisp, 1962; Clare et a/., 1985). The synchrony of 
spring bloom and egg hatching could be related to a component in the nutrition of 
adult barnacles. However, Crisp and Spenser (1958) showed that seawater extracts of 
unfed and fed adults were equally effective in inducing hatching. They proposed a 
barnacle hatching substance, endogenously produced by adults, and showed that the 
substance acts upon the musculature of mature embryos, not on the egg case. 

The hatching substance appeared to be a PG (Clare et a/., 1982, 1985). The sub- 
stance is extractable in a system optimized for PGs, it behaves like a PG on thin layer 
chromatography, and extracts of the dried cortex of a commercial source of PG (the 
gorgonian Plexura homomalla) acted biologically and chemically like barnacle hatch- 
ing substance. Extracts made in the presence of aspirin a PG-synthetase inhibitor 
in mammals did not induce hatching. Clare et al (1985) concluded that barnacle 
hatching substance is either a PG or a PG-like compound. 

Subsequent work underscores the importance of rigorous chemical methodolo- 
gies in indentification of biologically active compounds. Holland et al (1985) ex- 
tracted 50 kg of barnacles, then processed the extracts through four sequential systems 
of thin layer chromatography. The active compound was detected by bioassay at 
each stage. The purified compound was derivatized for gas chromatography-mass 
spectroscopy (GC-MS), which yielded a single major GC peak. Mass spectra of de- 
rivatized hatching factor and hydrogenated derivatized hatching factor were consis- 
tent, not with a PG, but with another eicosanoid, 10,11,1 2-trihydroxy-5,8, 11,17 eico- 
satetraenoic acid (Fig. 5). This compound is probably a lipoxygenase derivative of 
C20:5n3, an abundantly available fatty acid in marine invertebrates and also the pre- 
cursor of the 3-series PGs. 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 101 

OH OH 

OH 

FIGURE 5. Structure of barnacle hatching factor, 10, 1 l,12-trihydroxy-5,8,l 1,17-eicosatetraenoic 
acid. 



Reproduction in Mollusca 

PGs appear to stimulate egg production in the freshwater snail Helisoma durgi 
(Kunigelis and Saleuddin, 1986). When injected directly into the haemocoel of 
adults, ng quantities of PGE 2 produced apparent discomfort in all individuals and 
even death in isolated cases with no increase in egg masses or in eggs per mass. But 
when introduced into the female genital opening in a viscous fluid designed to ap- 
proximate semem, PGE 2 treatments stimulated a long-term increase in egg produc- 
tion. Four weeks after treatment of virgin snails with 25, 50, and 100 ng doses of 
PGE 2 , cumulative egg production was about 200, 425, and 650 eggs per animal, re- 
spectively. 

Reproductive tissues from virgin and mated snails, the ovotestis, seminal vesicle, 
bursa copulatrix, and oothecal gland presented substantial PG-biosynthetic activity 
in vitro. Mating significantly altered the activity in two of the tissues. In ovotestis, 
synthesis of PGE 2 decreased while PGA 2 synthesis increased with no change in syn- 
thesis of PGF 2(V . Synthetic activity changed in the bursa copulatrix, with PGE 2 and 
PGA 2 reduced to effective zero after mating; PGF 2(V was again unchanged. Differences 
in PG-synthetic activity did not occur in seminal vesicle or oothecal gland (Kunigelis 
and Saleuddin, 1986). These two lines of evidence the effects of PG treatments on 
egg production and alterations in PG-synthetic activity suggest that PGs play im- 
portant reproductive roles in this snail. 

PGs are also produced by accessory sex glands of another snail, Lymnaea stag- 
nalis (Clare et ai, 1986). Homogenates of the albumen gland, bursa copulatrix, pros- 
trate gland, and seminal vesicles converted radioactive arachidonic acid into labelled 
products that co-eluted with 6-keto-PGE,, PGE 2 , PGA 2 /B 2 (not resolved), throm- 
boxane B 2 (TxB 2 ), and several unknown compounds. Whole organs also converted 
arachidonic acid into these compounds, although in proportions different from the 
homogenates of the same organs. Effects of mating on PG-biosynthetic activity were 
not tested, nor were effects of PG administration on reproductive functions; nonethe- 
less, the PGs formed in the reproductive organs eventually may be shown to play a 
still undefined role. 

PGs induce spawning in two other molluscs, the abalone Haliotis refescens and 
the mussel Mytilus califorianus. When added to seawater cultures at 3 X 10" 12 M, 
PGE induced about a third and PGF about a half of male and female abalone to 
spawn (Morse et ai, 1977). Although the physiological mechanisms remain unclear, 
important biochemical insights have emerged. Addition of hydrogen peroxide to sea- 
water tanks induced synchronous spawning in H. refescens and M. califorianus. This 
observation is connected to the biochemistry of PG biosynthesis as understood in 
mammals. The first step in the conversion of arachidonic acid to the 2-series PGs is 
catalyzed by fatty acid cyclooxygenase (also known as prostaglandin endoperoxide 
synthetase). This involves first activation of the enzyme by a hydroperoxy group, then 
elimination of a hydrogen atom from C13 of arachidonic acid, leaving a free radical. 
This is followed by adding a peroxy radical in a bridge across C9 and Cl 1 , formation 
of the 8, 12 carbon-carbon bond (required for the cyclopentane ring in the final prod- 



102 D. W. STANLEY-SAMUELSON 

uct), isomerization of the 11,12 double bond to 1 2, 1 3, and addition of another peroxy 
radical to C15, with concomitant isomerization of the 12,13 double bond to 13,14. 
These final electron shifts generate PGG 2 , a short-lived intermediate in the conver- 
sion of an acid to PG. The hydrogen peroxide effect is pH dependent, with 
lower cone is releasing spawning at higher alkalinity. Morse et al. ( 1 977) sug- 
gested alkaline conditions (pH 9. 1 ) favored decomposition of hydrogen per- 
oxide highly reactive hydroperoxy free radical. Since a hydroperoxy group 
enzyme and peroxy radicals are added in two steps in the formation of 
PGG- . the free radicals derived from hydrogen peroxide may enhance overall conver- 
sion of precursor fatty acids to PGs. 

PGs appear to be important in basic physiological functions in molluscs, includ- 
ing ion regulation, possible renal function, and reproductive biology. This prelimi- 
nary work sets the stage for important questions of the precise physiological activity, 
and offers the possibility of gaining greater understanding of invertebrate physiology 
and appreciation of PGs in these systems. 

Oocyte maturation in starfish 

Starfish oocytes develop to the first meiotic prophase, then await the spawning 
period. Maturation, or meiosis reinitiation, is induced by a hormone produced and 
released by the follicle cells surrounding the oocytes, 1-methyladenine. Once stimu- 
lated by the hormone, the oocytes complete the developmental path leading to fertil- 
izable cells. 

Arachidonic and eicosapentaenoic acids also induce oocyte maturation in three 
species of starfish: Asterias rubens, Marthosterius glacialis, and Luidia ciliaris 
(Meijer et al., 1984). The PUFA-induced maturation is specific to these two fatty 
acids because 35 other fatty acids, ranging from C4:0 to C24:l and including satu- 
rated, monounsaturated, and polyunsaturated fatty acids, did not induce maturation. 
The maturation effect is dependent upon extracellular calcium and occurs at physio- 
logical concentrations (i.e., 50% maturation dose = 0.65 yuM arachidonic acid). The 
fatty acids stimulate the complete maturation program, including germinal vesicle 
breakdown, fertilization, and development into normal larvae. Fatty acids endoge- 
nous to the oocytes are able to stimulate maturation because addition of phospholi- 
pase A 2 , an enzyme that hydrolyses the fatty acid from the beta-carbon of phospholip- 
ids, also stimulated maturation. The phospholipase effect was calcium-dependent, 
and specific because phospholipases C and D did not bring on maturation. 

The hormone effect probably proceeds through release and metabolism of PU- 
FAs. Two phospholipase A 2 inhibitors in mammals, quinacrine and bromophenacyl 
bromide, inhibit hormone-stimulated maturation, which can be overcome by in- 
creasing 1 -methyl adenine concentrations. Five PGs did not stimulate maturation, 
and three cyclooxygenase inhibitors acetylsalicylic acid, indomethacin, and tolazo- 
line did not inhibit maturation. On the other hand, three lipoxygenase inhibitors 
in mammals quercetin, eicosatetraynoic acid and butylated hydroxytoluene did 
inhibit hormone-induced maturation. Four products of lipoxygenase metabolism of 
arachidonic acid, 12- and 1 5-hydroxyeicosatetranoic acids (HETE) and their corre- 
sponding hydroperoxyeicosatetraenoic acids (HPETE) stimulated maturation. 

Oocytes convert radioactive arachidonic acid into HETEs (Meijer et al., 1986a). 
Conversion of arachidonic acid does not occur in the absence of calcium, nor are 
oocytes stimulated to maturation. Following incubation with radioactive arachidonic 
acid, fractions with chromatographic behavior of HETEs were recovered and found 
to stimulate oocyte maturation. The lipoxygenase inhibitor eicosatetraynoic acid in- 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 103 

hibited both conversion of arachidonic acid and stimulation of oocytes. It would 
appear, then, that 1-methyladenine acts by release of PUFA, followed by conversion 
to a biologically active HETE, which induces maturation of the oocytes. 

Injection studies suggested that 12- and 15-HETE and corresponding HPETEs 
stimulated oocyte maturation (Meijer et a/., 1984). Upon re-evaluation, it was found 
that the tested compounds were contaminated with 5% of 8-HETE, the active com- 
pound in maturation (Meijer et al., 1986a). Meijer et al. (1987) showed that (8R)- 
HETE, but not (8S)-HETE, is produced by starfish oocytes. The R isomer is the only 
active compound when tested in pure form, and other lipoxygenase products, includ- 
ing other HETEs and leukotrienes are not active. 

A survey of eight starfish species shows that while the hormone 1-methyladenine 
stimulates maturation in all species, the stimulatory effect of arachidonic acid and 8- 
HETE occurs in only three of them (Meijer et al., 1986b). Species differences in re- 
sponse to various eicosanoids also have been observed in various physiological set- 
tings in mammals. At this early period of appreciating the possible physiological ac- 
tivities of these compounds in invertebrates systems, species differences underscore 
the hazards inherent in forming generalizations. 

Cercarial penetration of skin 

Eggs of the blood fluke Schistosoma mansoni leave their mammalian hosts in 
urine or feces, and continue larval development in snails. Free-swimming larvae 
called cercariae reinfect mammalian hosts by burrowing through the skin or by inges- 
tion with drinking water (Storer and Usinger, 1965). It has been known for a number 
of years that skin surface lipids stimulate cercarial penetration of animal membranes 
(Stirewalt, 1971). Among the skin surface lipids, free fatty acids, especially polyunsat- 
urated fatty acids, appeared to be most efficacious in stimulating penetration (Austin 
et al., 1972). Salafsky et al. ( 1984a) looked at the effect of certain fatty acids on two 
cercarial behaviors in vitro, namely cessation of swimming and initiation of penetra- 
tion. Their results show that certain PUFAs attracted cercariae to the center of their 
test membranes while monounsaturated fatty acids did not. A few fatty acids gave 
intermediate results because two monounsaturated fatty acids were as stimulatory as 
the PUFAs, and two other monounsaturates were less stimulatory than the PUFAs 
but were clearly more stimulatory than controls. Cyclooxygenase metabolites, rather 
than the PUFAs per se, may alter cercarial behavior. Two inhibitors of cyclooxygen- 
ase ibuprofen and, to a lesser degree, aspirin inhibited cercarial response to 
PUFA. 1 3-Azaprostanoic acid, thought to specifically antagonize the platelet throm- 
boxane/endoperoxide receptor in mammals, was also inhibitory. 

PUFAs and certain of their metabolites may affect cercarial penetration as well as 
modify behaviors that precede penetration. When Salafsky et al. (1984b) compared 
cercarial penetration into skin membranes prepared from essential fatty acid (EFA) 
deficient and EFA replete adult rats, they found about three times less penetration in 
the preparations from EFA deficient rats. Again, the inhibition may be related to 
formation of eicosanoids. Interperitoneal injections of ibuprofen led to a time-depen- 
dent accumulation of the drug in the skin of EFA replete rats. Cercarial penetration of 
the drug-treated skin was reduced. The percent inhibition increased with increasing 
amount of ibuprofen accumulated in the skin, up to a maximum inhibition of 
about 84%. 

When cercariae were incubated with radioactive linoleic acid, radioactivity could 
be recovered in high-pressure liquid chromatography fractions that eluted with 
PGE 2 , PGD 2 , LTC 4 , LTB 4 and 5-HETE. These data suggest that cyclooxygenase and 



104 D. W. STANLEY-SAMUELSON 

lipoxygenase sy unction within the cercariae. Radioimmunoassays of extracts 

from cercarf; oated with linoleic acid were also consistent with these products. 

Fusco et al. concluded that formation of eicosanoids is an essential step in 

penetn '^kin by cercariae of Schistosoma mansoni. If this can be sup- 

ported 1 'ork, it may present a rather interesting situation in which the 

P 7 e provided by a vertebrate host is metabolized into biologically active 

a parasite. 

yet known how the eicosanoids alter the behavior of the cercariae or 
enetration of mammalian skin. Fusco et al. (1985) suggest that vasodila- 
nch is induced by certain PGs, may help the parasite find and infiltrate the 
,/d system. It would appear that the eicosanoids, in this mode, would be usurped 
by the parasites to alter the host physiology. In this case, the finding by Rumjanek 
and Simpson ( 1 980) that adult worms do not synthesize PGE or PGF may be appreci- 
ated in terms of host physiology. On the other hand, the behavioral effects of cessation 
of swimming and initiation of penetration (Salafsky et al., 1984a), also induced by 
skin lipids, suggest a direct effect on the cercariae. 

Sponge cell aggregation 

Rich et al. (1984) suggest that the calcium dependent aggregation of marine 
sponge cells of Microcione prolifera is stimulated by leukotriene B 4 (LTB 4 ). LTB 4 
induced rapid cell aggregation in a dose-dependent way at 0.2 and 1 .2 ^M treatments. 
The effect appears to be specific for LTB 4 because eight PGs of A, B, D, E, and F 
series and eight lipoxygenase products failed to induce aggregation. 

The calcium ionophore A23187 and the species-specific aggregation factor 
(MAF) stimulate cell aggregation. The aggregating effects of these compounds can 
be inhibited by cyclooxygenase inhibitors including nordihydroquaiaretic acid and 
indomethicin, which also interfere with calcium flux. These data show that those 
agents which inhibit calcium flux also inhibit aggregation while those that promote 
calcium movement also promote aggregation. Interpretation is difficult because while 
a specific lipoxygenase product promotes aggregation, inhibitors of cyclooxygenase 
metabolism inhibit it. Perhaps both pathways are involved in cell aggregation, with 
LTB 4 stimulating PG formation, which then acts in concert with the LTB 4 . 

Egg-laying behavior in crickets 

The roles of PGs in insect reproduction were reviewed by Stanley-Samuelson and 
Loher (1986), from which the following summary is drawn. PGs were detected in 
extracts of various tissues from over a dozen species of insects. The most well under- 
stood physiological role of PGs is releasing egg-laying behavior in the field cricket 
Teleogryllus commodus. Adult females undergo sexual maturation, during which the 
abdomen becomes filled with hundreds of mature eggs. Certain behaviors that are 
likely to bring females into contact with males also develop. Insemination is achieved 
by transfer of a spermatophore to the genital organ of a female from where its contents 
migrate into the female's spermathecae. Cyclooxygenase activity is associated with 
the spermatophore contents, and once in the spermathecae of newly mated females, 
arachidonic acid is converted into PG. 

It is not known how the PG formed in the spermatheca releases egg-laying behav- 
ior, but increases in spermathecal and hemolymph PG titer after mating suggest that 
the PG acts at some site distant from the source. The observations that PGE 2 does 
not stimulate contraction of oviduct muscles in T. commodus (Loher, 1984) nor in a 



PHYSIOLOGICAL ROLES OF PROSTAGLANDINS 105 

cockroach (Cook et a/., 1984) and that oviposition behavior is a complex activity 
directed by the central, rather than peripheral, nervous system (Loher, 1984) support 
the hypothesis that the PGs function at the level of the central nervous system. 

Using egg-laying to assay structure-function relationships among a range of eico- 
sanoids, Stanley-Samuelson et al. (1986) found that highest egg-laying activity was 
associated with E-series PGs. The A-, B-, D- and F-series induced zero to intermediate 
egg-laying. Structures that departed from the basic PG structure, represented by 15- 
HETE and prostacyclin, were inactive. The 2-series PGs were more active than their 
1 -series analogues; hence, there may be a biological specificity for PGE 2 in releasing 
egg-laying behavior in that particular cricket species. 

Highest egg-laying activity was induced by 15-keto-PGE 2 . In mammalian sys- 
tems, this compound is formed by the action of prostaglandin dehydrogenase, located 
mainly in lungs, but also in liver and kidney. Biologically active PGE is rapidly 
cleared from the circulation of mammals by the activity of this enzyme. The observa- 
tion that a biologically inactive compound, in the usual mammal assays, was associ- 
ated with the greatest increase in egg-laying behavior marks a potentially important 
point in comparative physiology. Several features of the biology of eicosanoids appear 
to uniformly occur in the vertebrate and invertebrate systems as understood to date. 
For example, many compounds that inhibit the action of cyclooxygenase in mam- 
mals similarly inhibit the activity in invertebrates. On the other hand, as shown here, 
while the mammalian background will be important and useful in work on inverte- 
brate systems, fundamental differences are to be expected. 

PERSPECTIVES 

Various eicosanoids appear to be involved in the regulation of a variety of physio- 
logical and behavioral areas in representatives of many invertebrate phyla. In some 
cases (such as mediation of behavioral thermoregulation), the evidence for an eicosa- 
noid function is based on treatment of animals with a single compound and observa- 
tion of the response. At this level of observation, it remains to be established that 
eicosanoids are physiologically involved. Given a good base of preliminary observa- 
tions, important research goals would be to firmly show that, in the case at hand, PGs 
do mediate thermoregulatory behavior. In still other cases, such as the role of PG in 
releasing egg-laying behavior in crickets, there is sufficient evidence to accept that 
certain PGs do release egg-laying, although some details of the physiological mecha- 
nism where in the central nervous system PGs act and how they alter behavior 
are not yet understood. Research in this area could usefully be aimed, not at re- 
affirming the role of the eicosanoid, but at aquiring more details of the action. In study 
areas where considerable biochemical details are established as in starfish oocyte 
maturation cellular events remain unknown. Again, understanding how eicosa- 
noids act remains a major research goal. 

We are aware of eicosanoid roles in particular physiological areas in a given inver- 
tebrate organism. We know, for example, that PG releases egg-laying behavior in 
females of the cricket T. commodus. PGs are also detected in salivary glands, endo- 
crine glands, Malpighian tubules, testes, and ventral nerve cords. Aside from the 
known role in altering behavior, what do PGs contribute to the other tissue systems 
in which they appear? Are they involved in regulating ion flux in Malpighian tubules, 
secretion in salivary glands, and neural function in the nerve cord, within the same 
organism? 

Eicosanoids appear to be produced and to act at local, tissue, or cellular levels in 
mammals. PGE, produced by adipocytes functions within the same cells to modulate 



106 D. W. STANLEY-SAMUELSON 

the lipid mobilizing effect of certain hormones. Moreover, there are mechanisms that 

block PG circulation. The global circulation of PGE 2 , for example, is checked by the 

action of pro Jin dehydrogenase, located mainly in lungs, which converts the 

active com into a biologically inactive product. However, in the cricket T. 

comn ! dated that the release of egg-laying behavior by PGE 2 is mediated 

in i. >ike a broadly circulating hormone (Stanley-Samuelson and Loher, 

198' -Samuelson et ai, 1986). This point can be extended to research in the 

iction, at the whole-organism level, of eicosanoids in invertebrates. The 

xyeicosatetraenoic acid that functions as hatching substance in the barnacle 

oe an example of a compound produced in one organ system with its action 

observed elsewhere, again suggesting hormone action. 

With several likely roles of eicosanoids set forth, general research areas include 
establishing more firmly the activities, elucidating cellular details, and appreciating 
the possible modes of action of these compounds. One can assume that as details of 
eicosanoid action become known they will contribute greatly to our understanding 
of invertebrate physiology. 

ACKNOWLEDGMENTS 

I am grateful to Drs. W. Loher, R. H. Dadd, M. O. Theisen, and R. A. Jurenka 
for reading and making useful comments on the paper. The author and the work on 
T. commodus were supported by NIH grant RO 1 HD036 1 9 to W. Loher. 

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INTERSPECIFIC AGGRESSIVE BEHAVIOR OF THE 

LIMORPHARIAN CORYNACTIS CALIFORNICA 
vIDARIA: ANTHOZOA): EFFECTS ON SYMPATRIC 
CORALS AND SEA ANEMONES 

NANETTE E. CHADWICK 

Department of Zoology, University of California, Berkeley, California 94720 

ABSTRACT 

Corallimorpharians are sessile cnidarians that are morphologically similar to the 
actiniarian sea anemones and scleractinian corals. This study describes for the first 
time the behavioral mechanism and effects of aggression by a corallimorpharian. Pol- 
yps of the temperate clonal corallimorpharian Corynactis californica extruded their 
mesenteries and associated filaments onto members of certain species of sea anemo- 
nes and corals. They did not exhibit this behavior intraspecifically, and members 
of different clones of C. californica remained expanded upon contact. In contrast, 
members of four species of corals and zoanthids responded to contact with C. califor- 
nica by contracting their tentacles, and members of three sea anemone species bent 
or moved away, detached from the substrate, or attacked using their aggressive struc- 
tures. When interspecific contact was prolonged, individuals of C. californica ex- 
truded filaments onto, and killed polyps of, the sea anemones Anthopleura elegantis- 
sima and Metridium senile within 3 weeks, and the corals Astrangia lajollaensis and 
Balanophyllia elegans within 4-10 months under laboratory conditions. The use of 
extruded mesenterial filaments by C. californica to attack members of other antho- 
zoan species is similar to the aggressive behavior exhibited by many scleractinian reef 
corals. Field observations suggest that C. californica may use this agonistic behavior 
during interspecific competition for space on hard marine substrate. 

INTRODUCTION 

Some of the most striking behaviors exhibited by members of the class Anthozoa 
(Phylum Cnidaria) are the aggressive behaviors of certain actiniarian sea anemones 
and scleractinian corals. Corals may attack competitors using sweeper tentacles 
(Richardson et al., 1979; Wellington, 1980; Bak et ai, 1982; Chornesky, 1983; Hi- 
daka and Yamazato, 1984), sweeper polyps (Sheppard, 1982), extruded mesenterial 
filaments (Lang, 1973;Glynn, 1974;Loya, 1976; Cope, 1981; Bak?/ al., 1982; Logan, 
1984), or nematocysts discharged from the colony surface (Rinkevich and Loya, 
1983), and actiniarian sea anemones may use elongated catch tentacles (Williams, 
1975; Purcell, 1977) or marginal vesicles called acrorhagi (Bonnin, 1964; Francis, 
1973b;Ottoway, 1978; Bigger. 1980; Brace, 1981;Ayre, 1982;Sebens, 1984). These 
aggressive responses are complex. They often involve the induced morphogenesis and 
directed application of specialized structures packed with nematocysts (Purcell, 1 977; 
Chornesky, 1983; Watson and Mariscal, 1983; Hidaka and Yamazato, 1984;Hidaka, 
1985), and may be initiated upon recognition of other genotypes or species of antho- 
zoans(Lang, 1973; Bigger, 1980). 

Received 2 March 1987; accepted 20 May 1987. 

110 



CORALLIMORPHARIAN BEHAVIOR 1 1 1 

However, little is known about the aggressive behavior of another group of antho- 
zoans, the corallimorpharians. Sebens (1976) reported the effects of competitive in- 
teractions between corallimorpharians and other anthozoans on the Caribbean coast 
of Panama, but did not specify the behaviors they used. The only other study relating 
to corallimorpharian behavior is that of Hamner and Dunn (1980), who described 
the unique feeding mechanism of some tropical Pacific corallimorpharians in which 
prey are enfolded in the oral disk. 

Corallimorpharians occur throughout the world (Carlgren, 1949) and may be 
abundant on temperate rocky shores (Hand, 1955; Forster, 1958; Pequegnat, 1964; 
Castric-Fey el a!., 1978; Foster and Schiel, 1985), as well as on tropical coral reefs 
(Fishelson, 1970; den Hartog, 1980). Certain members of this group form clonal ag- 
gregations that cover large areas of hard substrate, and are the dominant sessile organ- 
isms in some temperate marine communities (Forster, 1958; Castric-Fey el ai, 1978). 
Thus, interactions of corallimorpharians with other sessile organisms may have im- 
portant consequences for the structure of these communities. 

Corallimorpharians superficially resemble the actiniarian sea anemones in that 
they lack a calcareous skeleton (Carlgren, 1949). However, they are more like the 
stony corals in most other aspects of their morphology: they lack basilar muscles, may 
have tissue connections between adult polyps, lack ciliated tracts on their mesenterial 
filaments, and their cnidae are similar to those of corals (Carlgren, 1949; Schmidt, 
1 974; den Hartog, 1 980). In light of the morphological relationships among members 
of these three anthozoan groups, a comparison of their aggressive behaviors is of 
interest. 

This study describes the interspecific aggressive behavior of the temperate clonal 
corallimorpharian Corynactis californica. This behavior was first recorded in Chao's 
(1975) unpublished student paper. He observed that C. californica extruded mesen- 
terial filaments to damage the sea anemones, 4 nthopleura elegantissima and Melrid- 
ium senile during interspecific interactions in the laboratory. This is aggression, 
which is defined by Webster's Third New International Dictionary as "an offensive 
action or attack," and in this instance is elicited upon contact with the anemones. 
Haderlie el al. (1980) briefly mentioned this behavior in their account of the natural 
history of C. californica. The present paper expands on these reports by presenting a 
quantitative analysis of mesenterial filament extrusion by C. californica, the specific- 
ity of this aggressive response, and its effect on the behavior and survival of some 
common sea anemones and corals under laboratory conditions. 

Natural history 

Corynactis californica Carlgren 1936 is the only species of corallimorpharian to 
occur along the west coast of North America, where it ranges from Washington State 
(Birkeland, 1971) to San Benitos Island in central Baja California (J. Engel, Tatman 
Foundation, pers. comm.). Members of this species reproduce asexually by fission 
(Hand, 1955) and budding (pers. obs.) to form aggregations on hard substrate (Fager, 
1971; Haderlie et al., 1980; Foster and Schiel, 1985), from the lower intertidal zone 
(Hand, 1955) to at least 50 meters depth (Birkeland, 1971; Schmieder, 1984, 1985). 
C. californica polyps are common on the vertical faces of subtidal rock reefs where 
they attain densities of up to 3000 polyps per square meter (Pequegnat, 1964). In 
southern California, field experiments show that the presence of aggregations of C. 
californica may increase the abundance of rock oysters (Vance, 1978) and mussels 
(Landenberger, 1967; Wolfson et ai, 1979) by protecting them from predation by sea 
stars. Groups of this corallimorpharian form interspecific boundaries with clones of 



112 N. E. CHADWICK 

the sea anemones Anlhopieiira elegantissima and Metridium senile on artificial sub- 
strates such as wharf pilings (Chao, 1975; Haderlie and Donat, 1978) and offshore oil 
platforms (Carlisle el al, 1964). Groups of C californica also co-occur on subtidal 
rock reefs in ke >is with several species of corals, most commonly the colonial 

coral As i -ant ;<v/.s7.v (Pequegnat, 1964) and the solitary corals Balanophyllia 

elegcu; nhus stearnsii (Pearse and Lowry, 1974; Lewbel el al. , 1981; Fos- 

, 1985; North et al., 1985). 

: cs greatly between different clonal aggregations of C californica. Clones 
rn. pink, orange, or occasionally blue or purple. Members of each aggrega- 

tion asexually produce polyps of the same color in both the laboratory (pers. obs.) 
and the field (Turner et a I., 1 969). Thus, color in this species appears to be genetically 
controlled, and in the present study polyps from different, distinctively colored aggre- 
gations were assumed to be genetically different (non-clonemates). 

MATERIALS AND METHODS 
Collection and maintenance of organisms 

Specimens of C. californica and the other organisms used in this study were col- 
lected at four sites along the coast of central California (Table I). Laboratory experi- 
ments were conducted between June 1984 and July 1986 at three facilities of the 
University of California: Bodega Marine Laboratory. Joseph M. Long Marine Labo- 
ratory, and in cold rooms on the Berkeley campus. Organisms were maintained in 
plastic trays supplied with flowing seawater at ambient sea temperature (13-15C), 
or in closed refrigerated aquaria filled with aerated natural seawater. All tanks were 
cleaned and animals fed adult brine shrimp (Anemia salina) weekly. 

Mechanism and specificity of aggressive behavior 

The first set of experiments focused on a description of mesenterial filament extru- 
sion by C californica, and determination of the stimuli that elicit this response. Only 
fully expanded, undamaged individuals of C. californica were used, and all within 
two weeks of collection. Polyps were brought into contact with a range of physical 
and biological stimuli (Table I) to elicit extrusion. Polyps were observed continuously 
for the first hour of contact, then once each hour for at least 12 h, and then intermit- 
tently for several days. A different individual of C. californica was used for each obser- 
vation; Table I shows the number of replicate observations with each stimulus. Data 
were collected on the diameter and behavior of each polyp, occurrence of mesenterial 
filament extrusion, interval from the start of contact to extrusion, duration of extru- 
sion, and the origin and maximal length of any extruded filaments. 

Effects ofC. californica on selected anthozoans 

During the above contacts between polyps of C. californica and seven other spe- 
cies of anthozoans (Table I), data were also collected on the behavioral response of 
each anthozoan. Their responses to C. californica were categorized as: contracted, 
expanded, bent the column away, moved away on the pedal disk, detached from the 
substrate, or attacked C. californica. During trials between C. californica and mobile 
anthozoans such as the actiniarian sea anemones, the latter were repeatedly moved 
back into contact with C. californica to allow adequate time for a response. 

The second set of experiments examined effects of C. californica on the survival 
of selected anthozoans over several weeks in the laboratory. To test the effect of C. 



CORALLIMORPHARIAN BEHAVIOR 113 

californica on actiniarian sea anemones, individuals of the clonal anemones An- 
ihopleura elegant issima and Mctridium senile were placed in the center of groups of 
C. californica that were attached to shells or rocks. This method prevented movement 
away from contact by the anemones. Control anemones were placed on rocks that 
were interspersed in the same tray with experimental groups, but not in contact with 
C. californica. Data were then collected on the behavior and condition of the anemo- 
nes once a week for three weeks. 

Effects of prolonged contact with C. californica also were examined in two species 
of scleractinian corals, Astrangia lajollaensis and Balanophyllia elegans. Individual 
corals were attached to glass microscope slides or shells using H. A. Calahan's Ma- 
rinepoxy (Davis Instruments, San Leandro, CA 94578). This epoxy has been used on 
anthozoans for several years in the laboratory without apparent harm (J. S. Pearse, 
University of California, Santa Cruz, pers. comm.). Barnacle shells bearing aggrega- 
tions of C. californica were broken into small bits, and each piece of shell bearing a 
single polyp of C. californica was cemented adjacent to a coral. Empty shells with no 
C. californica were glued next to other corals as controls. Experimental and control 
plates of corals were then intermingled in trays of seawater, and the condition and 
behavior of each polyp was recorded once each month for 1 2 months. During this 
time, polyps of the asexually reproducing species budded off new individuals, and 
each month these were counted and the degree to which they had overgrown other 
polyps was determined. 

RESULTS 
Description of aggressive behavior in C. californica 

Upon contact, the tentacles of individual C californica adhered to those of polyps 
of certain other anthozoans. Then the interacting polyps often contracted slightly and 
their tentacles retracted. Over the next few minutes, the two polyps went through 
several cycles of expansion, contact, contraction, and re-expansion. If they main- 
tained fairly constant tentacular contact, a mass of highly convoluted mesenteries 
and their associated filaments eventually appeared at the mouth or through a break 
in the body wall of the C. californica polyp (Fig. la). These filaments were withdrawn 
into the coelenteron at the end of each extrusion. Of 214 C. californica individuals 
observed, most extruded filaments through the mouth (69%), through openings in 
the column (7%), or along the junction of column and base (24%) of the polyp. One 
polyp put out filaments through the tips of its tentacles. These openings in the body 
wall were temporary and healed soon after the mesenterial filaments were withdrawn. 

C. californica individuals almost always directed filaments laterally toward the 
side on which they had been stimulated (in 98% of cases, n == 214). These filaments 
then adhered to the source of stimulation and spread over its surface. They appeared 
highly extensible (Fig. Ib), and if the stimulus source was pulled away, the filaments 
could be stretched up to four times the diameter of the polyp to which they belonged. 
Extruded filaments ranged in length from 1 to 42 mm (median == 3 mm). However, 
most polyps extruded filaments only 1-10 mm in length (91% of polyps, n = : 190), 
or about 0.1-1.5 times polyp diameter. Extrusion length did not vary with polyp size; 
small (5 mm diameter) polyps often extruded filaments at least 10 mm in length, 
while many large (>15 mm diameter) polyps put out filaments only 2-4 mm long. 
Often several mesenteries with their attached filaments were extruded by a single 
polyp, and they frequently spread to cover the organism that was the source of stimu- 
lation. 



114 



N. E. CHADWICK 




FIGURE 1 . A. Side view of extrusion of mesenterial filaments by an individual of the corallimorphar- 
ian Corynactis californica onto a contracted polyp of the actiniarian sea anemone Metridium senile (left). 
B. Top view of mesenterial filament extrusion by two polyps of C. californica (center) onto a retreating 
individual of the actiniarian sea anemone Anthopleura elegantissima (upper right). Photo by Galen Rowell. 
Note that in both photographs the filaments extend toward the actiniarians, and that in B they adhere to 
the anemone as it moves away. Scale bars = 1 cm. 



CORALLIMORPHARIAN BEHAVIOR 



115 



30r 



(fl 
<a 

g 
'> 

TO 



Q) 
O 

<D 
CL 




10 - 



time (hours) 

FIGURE 2. Time from start of contact to start of mesenterial filament extrusion by individuals of 
Corynactis californica upon contact with members of other anthozoan species and large food items (n 
= 190, median = 2.5 h, range = 0.5-72 h). 



The interval from the start of contact to the start of extrusion ranged from 0.5 
to 72 hours, but most individuals began to extrude filaments within a few hours of 
application of an appropriate stimulus (Fig. 2). At least 0.5 h of continuous contact 
was necessary to elicit extrusion; when contact was intermittent, extrusion often be- 
gan only after several days. Most extruded filaments reached their maximal length 
1-12 h from the start of contact (median = 7 h, range = : 1-72 h, n = 129), and then 
were slowly withdrawn back into the coelenteron. The duration of extrusion varied 
greatly (median = 7 h, range =~- 1-144 h, n = 149); when contact with an appropriate 
stimulus was continuous, the filaments of some polyps remained extruded for up to 
six days. 



Specificity of filament extrusion by C. californica 

C. californica polyps extruded mesenterial filaments most frequently upon con- 
tact with certain types of biological stimuli (Table I). They did not respond to conspe- 
cifics, and instead, both clonemate and nonclonemate polyps remained expanded 
and intermingled their tentacles during contact. In contrast, a large percentage of C. 
californica individuals extruded mesenterial filaments onto members of three species 
of actiniarian sea anemones and the scleractinian coral Astrangia lajollaensis (Table 
I). Extrusion onto the solitary corals Paracyathus stearnsii and Balanophyllia elegans 
was less frequent and often occurred only after 12 or more hours of contact. All ten 
polyps of C californica that extruded filaments onto P. stearnsii did so 1 3-50 h from 
the start of contact, and extrusion was observed onto B. elegans only after several 
days or weeks from the start. Few C. californica individuals responded to the zoanthid 
Epizoanthus scotinus (Table I). 

C. californica polyps rarely used mesenterial filaments to attack other sessile or- 
ganisms such as hydroids, colonial tunicates, sponges, or algae (Table I). However, 
they did extrude filaments onto food items that were too large to ingest (Table I). To 
assess the size threshold for ingestion of large food items, expanded individuals of C. 
californica were offered pieces of fish that were less than, equal to, or slightly greater 
than their own polyp volume (by visual estimate). The polyps injested food items 
that were smaller than or equal to their own volume in 38/49 cases (78%). When 
offered larger prey, however, they almost always extruded filaments over the food 
(Table I). 



116 



N. E. CHADWICK 



TABLE I 

Collection sites, sti ' percent of Corynactis californica that extruded mesenterial 

filaments onto ea> ' .-rimulns 





% C. californica 


Collection 


that extruded 


site Stimulus 


Common name filaments 



ANTHOZOANS 
BCHM Corynactis californica 

clonemates 

non-clonemates 

MB Anlhopleura elegantissima 

MB Metridiwn senile 

B Epiactis prolifera 

H Astrangia lajollaensis 

CH Paracyatluts stearnsii 

H Balanophyllia elegans 

C Epiioanthns scot inns 

NON-ANTHOZOAN SESSILE ORGANISMS 

C A/lopora californica 

C Garveia annulata 

H Acarnns erilhicii.s 

H Diaperoecia californica 

H Archidistoma psammion 

H Cystodytes lobata 

H Rhodymenia pacifica 

LARGE FOOD ITEMS 
MB Mytilns ednlis 

H Sebasles spp. 

PHYSICAL STIMULI 

Puncture with a glass needle** 
Contact with a sterile glass rod 



Corallimorpharian 



Actiniarian sea anemone 
Actiniarian sea anemone 
Actiniarian sea anemone 
Scleractinian coral 
Scleractinian coral 
Scleractinian coral 
Zoanthid 



Hydrocoral 
Hydroid 
Sponge 
Bryozoan 
Colonial tunicate 
Colonial tunicate 
Red alga 



Bay mussel 
Rock fish 



(28) 

(40) 

97. 7* (44) 

89.4* (38) 

100* (17) 

88.9* (27) 

43.5 (23) 
6.5 (31) 

13.6 (22) 



23.5 




11.1 

4.8 
13.3 





(17) 
(10) 
(28) 
(18) 
(21) 
(15) 
(21) 



100* (21) 
83.0* (53) 



4.8 (21) 
2.4 (42) 



Numbers in parentheses indicate the number of polyps of C californica exposed to each stimulus. 

Collection sites: B = Breakwater at Doran Beach Park, Bodega Bay, Sonoma County, CA, on intertidal 
boulders, C = Cordell Bank, Marin County, CA, on rock pinnacles at 40-50 m depth, H = Hopkins Marine 
Life Refuge, Monterey County, CA, on rock reefs at 10 m depth, M = Monterey Municipal Wharf #2, 
Monterey County, CA, intertidaily on wharf pilings. 

* Responses significantly greater than those to all other stimuli, G-test for homogeneity of replicates, 
G= 14.08, P<. 05. 

** The column of each polyp of C. californica was punctured with a sterile glass needle, which was left 
in place for at least 1 2 h. 



Differences were observed in the quality of extrusion onto food items versus an- 
thozoans. When presented with large pieces of fish or mussel, most C. californica 
expanded, pressed their oral disks and tentacles onto the food, and extruded filaments 
out through their mouths (Table II). In contrast, when contacting anthozoans such 
as sea anemones or corals, C californica often contracted and/or put out filaments 
laterally through openings in the body wall (Table II). Filaments extruded onto an- 
thozoans also were significantly longer than those extruded onto prey items (Fig. 3). 

In response to physical contact with an inert glass rod, or physical damage to the 
column wall, C. californica rarely extruded filaments (Table I). However, polyps did 
put out filaments when subjected to extreme physical stress, such as when they were 
accidentally crushed or became desiccated. Individuals also occasionally extruded 
filaments in the absence of any apparent stimuli. 



CORALLIMORPHARIAN BEHAVIOR 



117 



TABLE II 

Comparison of behavioral responses to different stimuli (food items versus anthozoans) 
by Corynactis californica during extrusion ofmesenterial filaments 

Number of C. californica with each type of response 
during extrusion 



Posture: 


Expanded Contracted 


Filament 
Stimulus origin: 


Mouth 


Body wall Mouth 


Body wall 


Large food items (fish, mussel) 


63 


1 1 





Anthozoans 








(corals, zoanthids, sea anemones) 


24 


29 54 


23 



See Table 1 for species of stimuli used. 

The distribution of the responses is dependent upon the type of stimulus contacted (R X C test of 
independence using G-test, G = 62.61, P < .0 1 ). 



Effects of C. californica on the behavior and sunival of other anthozoans 

Contact with polyps of C. californica caused strong avoidance or attack responses 
by most of the anthozoans tested (Table III). However, conspecific C. californica of 
different genotypes (non-clonemates) did not avoid each other, and most remained 
expanded during contact. Non-mobile anthozoans of other species, such as sclerac- 
tinian corals and zoanthids, contracted their tentacles and often their entire polyps 
within minutes when placed in contact with C. californica (Table III). Two individu- 
als of the coral Paracyathus stearnsii extruded their mesenterial filaments at 7 h but 
these did not extend far enough to contact or damage C. californica polyps. The 
actiniarian sea anemones varied in response depending upon whether they were sur- 
rounded by C. californica polyps. When not surrounded, most individuals of An- 
thopleura elegantissima and Metridium senile bent away, moved away via pedal loco- 
motion, or attacked the corallimorpharian (Table III). Three polyps of A. elegantis- 
sima inflated their specialized aggressive structures called acrorhagi and applied them 
to C. californica at 0.5-2 h. These attacks left acrorhagial peels that caused localized 



V) 



.C 
<*- 

o 



40 



30 



20 



10 








123456789 10 y >10 

length of filaments extruded (mm) 

FIGURE 3. Comparison of length of mesenterial filaments extruded by Corynactis californica in re- 
sponse to large food items (shaded bars, n = 65, median = 2 mm, range =1-10 mm) versus anthozoans 
(striped bars, n = 123, median = 5 mm, range = 1-42 mm). A significant difference exists between the two 
populations (normal approximation to the Wilcoson rank sum test. Z = 6.83, P < .01). See Table I for 
species used. 



118 



N. E. CHADWICK 



TABLE III 

Variation in the bei responses of selected anthozoans to contact with polyps of the 

corallimorphar,a.- lactis californica 



Number with each behavioral response 


Total 
number 
p.nthozoan tested Expand Contract 


Bend 

away 


Move 
away 


Detach 

base 


Attack 



, CLONEMATE 
CONSPECIFICS 

OF C. californica 40 38 

CORALS 106 12 

ZOANTHIDS 20 

SEA ANEMONES 
Anthopleura 

elegantissima 

not surrounded 34 

surrounded 1 5 4 

Metridium senile 

not surrounded 22 

surrounded 18 3 

Epiactis prolifera 
not surrounded 14 



2 

92 
20 




6 

1 
1 











2 


2 


1 









29 


8 












5 

1 
5 





2 a 





3 h 


5 c + 5 d 
9 c 





Types of attack: a = extrusion of mesenterial filaments by the coral Paracyathus stearnsii at 7 h; b 
= acrorhagi; c = extruded acontia; d = catch tentacles, used by 5/10 individuals that possessed them. See 
Table I for species of corals and zoanthids used. 

Not surrounded/surrounded indicates whether or not each anemone was surrounded by polyps of C. 
californica during the interaction. 

The distribution of responses was dependent upon the type of anthozoan involved (R X C test of 
independence using G-test, P < .0 1 ). 



damage to C. californica polyps, but the damaged areas healed within a few days. 
Five out of ten polyps of Metridium senile that possessed well-developed aggressive 
structures (catch tentacles) also inflated and applied them to polyps of C. californica 
within 2-1 1 h of contact. However, none of these catch tentacles adhered to the coral- 
limorpharians, and they did not appear to cause damage. Metridium senile also fre- 
quently extruded acontia onto C. californica, both when surrounded and not sur- 
rounded (Table III). The acontia adhered strongly to and killed some C. californica 
individuals. Polyps of the actiniarian sea anemone Epiactis prolifera were tested only 
when not surrounded, and most avoided contact within 3 h by moving away on the 
substrate or detaching their pedal disks and then rolling or floating away (Table III). 

Individuals of the sea anemones M. senile and A. elegantissima were killed within 
one to three weeks (Fig. 4a) during prolonged contacts with surrounding groups of C. 
californica polyps. These anemones often detached from the substrate, but adhered to 
the tentacles of the surrounding C californica polyps and were unable to escape. They 
were then repeatedly attacked by the extruded mesenterial filaments of C. californica, 
and their tissues became necrotic within a few days. Control anemones that did not 
contact C. californica remained expanded and firmly attached to the substrate 
throughout the experiment. 

Corynactis californica had a much slower but fatal effect on members of two spe- 
cies of scleractinian corals. Within two weeks from initial contact, C. californica had 
caused tissue damage to most individuals of the corals Astrangia lajollensis (57/74 
polyps damaged, =77%), and Balanophyllia elegans (12/19 polyps damaged, =63%). 



CORALLIMORPHARIAN BEHAVIOR 



119 



A. sea anemones 



_a> 

!5 


o 



100- 



75- 



50- 



25- 




T 



T 



contact 
Corynactls 



no 
contact 



1 2 

time (weeks) 



-f- 100 1 
C 
0) 

o 

fe 7S 

a. 



50- 



25- 



J 



B. corals 




contact 
Corynactis 



no 
contact 



2468 

time (months) 



10 



12 



FIGURE 4. Effect of contact with the corallimorpharian Corynactis californica on the survival of 
selected sea anemones and corals. Bars represent 95% confidence limits. See text for details. A. Effect on 
polyps of the actiniarian sea anemones A nthopleura elegantissirna (solid lines, n = 21 contact, n = 20 no- 
contact) and Metridium senile (dashed lines, n = 17 contact, n = 17 no-contact). At three weeks, the 
proportions of anemones killed in the experimental (contact) versus control (no-contact) groups were sig- 
nificantly different for both species (G-test of independence for proportions, P < .0 1 ). B. Effect on polyps 
of the scleractinian corals Astrangia lajollaensis (solid lines, n = 69 contact, n = 3 1 no-contact) and Balano- 
phyllia elegans (dashed lines, n = 19 contact, n = 21 no-contact). At twelve months the proportions of 
corals killed in the experimental (contact) versus control (no-contact) groups were significantly different 
for both species (G-test of independence for proportions, P < .01 ). 



During the ensuing months, C. californica polyps asexually produced many new indi- 
viduals which eventually grew over and around the corals. In 6 months on one plate, 
10 C. californica individuals produced over 80 polyps that killed and covered the 
original 10 A lajollaensis polyps. After 12 months of contact, corallimorpharian pol- 
yps had killed most of the corals on the experimental plates (Fig. 4b). 



120 N. E. CHADWICK 

C. californica individuals appeared to affect only the corals that they touched. In 
several cases, tissue was damaged and calcareous skeleton was exposed only on the 
side of a coral that faced toward a C. californica polyp. In cases where asexually pro- 
duced po! alifornica grew away from and ceased to contact the experimental 
corals, the latter remained healthy and undamaged. Control corals that were isolated 
from contact with C. californica also remained alive (Fig. 4b), and during the year 
produced many new polyps, presumably both sexually (via brooded planulae) in Ba- 
lanophvllia elegans, and asexually (via clonal budding) in Astrangia lajollaensis. 

DISCUSSION 

This report is the first detailed description of aggressive behavior in a coralli- 
morpharian. The type of aggression exhibited by C californica, extrusion of mesente- 
rial filaments, is very similar to the attack behavior of many tropical scleractinian 
corals (Lang, 1973; Glynn, 1974; Loya, 1976; Wellington, 1980; Cope, 1981; Bak et 
ai, 1982; Logan, 1984). C. californica and certain corals readily extrude their mesen- 
terial filaments onto members of other anthozoan species and onto large food items 
(Table I; Yonge, 1930a; Lang, 1973). The timing of the extrusion response is also 
remarkably similar in corals and C. californica. Lang (1973) reported that Jamaican 
reef corals extruded their filaments 0.5-12 h after initiation of contact with certain 
coral species, and Glynn (1974) noticed extrusion by eastern Pacific corals 8-12 h 
after contact with competing corals. In the present study, most C. californica individ- 
uals also put out their filaments within 12 h (Fig. 2). The extruded filaments of both 
C. californica and scleractinian corals cause extensive damage to and eventually kill 
other anthozoans if contact is prolonged and if, in corals, the other colony is small 
enough (Lang, 1973; Fig 4). Since corals and corallimorpharians are morphologically 
very similar (den Hartog, 1 980), one might expect to see this similarity in their aggres- 
sive behaviors as well. This type of aggression, via mesenterial filament extrusion, 
differs from the competitive behavior of some of the actiniarian sea anemones that 
coexist with C. californica and use their marginal spherules (Francis, 1973b) or catch 
tentacles (Purcell, 1977) to attack competitors. These differences in behavior under- 
score the major morphological differences between a corallimorpharian such as 
C. californica, and actiniarian sea anemones. They also support the idea that coralli- 
morpharians are more closely related to scleractinian corals than they are to sea 
anemones. 

Unlike the specialized aggressive structures of actiniarian sea anemones that are 
used only during competitive interactions (Bonnin, 1964; Francis, 1973b; Williams, 
1975; Purcell, 1977; Watson and Mariscal, 1983), the mesenterial filaments of C 
californica appear to serve a variety of functions. In all anthozoans studied thus far, 
the mesenterial filaments are the major sites for digestion and absorption of food in 
the coelenteron (Yonge, 1930b; Nicol, 1959; Van-Praet, 1985). These filaments con- 
tain gland cells that secrete strong proteolytic enzymes, as well as nematocysts that 
may inject cytolytic toxins into prey (Van-Praet, 1985). Special areas on the filaments 
and adjacent mesenteries then absorb the partially digested foodstuffs (Yonge, 1 930b; 
Van-Praet, 1985). Corynactis californica also extrudes mesenterial filaments onto 
food that is too large to take into the coelenteron (Table I), presumably to digest it 
externally. This behavior allows polyps to consume a large range of prey sizes. Certain 
tropical Pacific corallimorpharians envelope prey in the oral disk, and then extrude 
filaments out of the mouth to digest them (Hamner and Dunn, 1980). Many species 
of reef-building corals also consume prey externally via extruded filaments (Carpen- 
ter, 1910; Yonge, 1930a, 1968; Goreau et ai, 1971). Thus, two major functions of 



CORALLIMORPHARIAN BEHAVIOR 121 

mesenterial filaments in these organisms appear to be the internal breakdown and 
absorption of food, and external consumption of large prey. Mesenterial filaments 
also are used during physical stress. Some corals extrude their filaments when oil is 
introduced into their coelenterons (Bak and Elgershuizen, 1976), when they are ex- 
posed to intense light (Lang, 1973), or when they are handled roughly (Duerden, 
1 902). C. californica polyps also exhibit extrusion when stressed (see Results). Finally, 
divers observed C californica polyps extruding their mesenterial filaments onto one 
of their major predators, the sea star Dermasterias imbricata (Annett and Pierotti, 
1984; pers. obs.). Thus, the lobed filaments along the edges of anthozoan mesenteries 
may serve multiple functions in certain corals and corallimorpharians. 

An interesting aspect of the aggressive/defensive use of mesenterial filaments by 
C. californica is the complete lack of response to conspecifics (Table I). Members of a 
given clonal aggregation presumably would benefit from damaging and overgrowing 
those of a different, genetically distinct aggregation (as discussed by Francis, 1973b). 
However, in the field distinctly colored groups of C. californica often intermingle and 
show no evidence of aggression or damage along their interacting borders (pers. obs.). 
The wide, anemone-free zones that are visible between aggregations in other species 
known to show interclonal aggression (Francis, 1973a; Purcell, 1977) do not occur 
in this species. Most reef corals that use mesenterial filaments to attack competitors 
also only extrude them interspecifically (Lang, 1973; Cope, 1981 ). One exception has 
been reported: the Caribbean coral Montastrea annularis appears to extrude fila- 
ments onto conspecific colonies to damage them (Logan 1984, 1986). 

The present study demonstrates that under laboratory conditions C. californica 
strongly affects both the behavior and survival of certain other anthozoans (Table III, 
Fig. 4). These results have several ecological implications. In shallow subtidal habitats 
along the coast of California where C. californica occurs, hard surfaces are often com- 
pletely covered with organisms (Pequegnat, 1964; Haderlie and Donat, 1978; Vance, 
1 978; Schmieder, 1 984, 1 985), and space for settlement and growth may be a limiting 
resource for sessile animals. The species of sea anemones tested in this study often 
moved away from or otherwise avoided contact with C. californica (Table III); where 
they co-occur in the field, this behavior might free space for growth along the interspe- 
cific borders of C. californica aggregations. The avoidance responses of these sea 
anemones are the same behaviors used to effectively escape attack by conspecifics 
(Francis, 1973b; Purcell, 1977) and predators (Waters, 1973; Edmunds el ai, 1976). 
However, the specialized aggressive structures ofAnthopleura elegantissima and Me- 
tridium senile apparently were not effective against C. californica (see Results). The 
acontia of M. senile caused the most damage to C. californica, and in the field may 
allow the former to kill polyps of the latter along their interacting borders. L. Harris 
(University of New Hampshire, pers. comm.) has observed that, in the laboratory, 
M. senile also uses acontia to attack individuals of the sea anemones A elegantissima. 
Actinia equina, and Urticina (= Tealia) piscivora. 

The results of the present study differ somewhat from those presented by Chao 
(1975). He described an aggressive hierarchy in which C. californica was dominant 
over A. elegantissima, while M. senile was dominant over both of the former species. 
The present results confirm that C. californica causes tissue damage to A. elegantis- 
sima (Table III, Fig. 4a), but show that C. californica and M. senile damage each 
other, with no clear competitive outcome. A clear dominance ranking of these three 
cnidarian species remains to be determined. 

Field observations also suggest that C. californica damages sea anemones and cor- 
als under natural conditions. Chao (1975) noticed that interspecific boundary areas 
about 2-3 cm wide occurred between aggregations of C. californica and the sea anem- 



122 N. E. CHADWICK 

ones Anthopleura elegantissima and Metridium senile found on intertidal pilings at 
the Monterey Wharf. Francis (1973b) and Purcell (1977) showed that anemone-free 
zones between :<ups within the latter two species are maintained by aggression be- 
tween cio^ orridors along their boundaries with C. californica could be main- 
tained by oidance behaviors of the anemones (Table III), or by the death of 
anemone 1 ;hat have been repeatedly attacked by the mesenterial filaments of C. cali- 
for> i-a). 

subtidal rock reefs, certain scleractinian corals also appear to be negatively 

cted by contact with C. californica. Fadlallah (1981) observed a polyp of C. califor- 
nica extruding filaments onto and killing an individual of the solitary coral Balano- 
phvllia elegans in the kelp forest at Hopkins Marine Life Refuge (HMLR) in Monte- 
rey. Polyps of B. elegans and the colonial coral Astrangia lajollaensis that occur adja- 
cent to C. californica on subtidal boulders at HMLR often show damaged tissues and 
exposed skeletons (pers. obs.). In addition, the vertical distribution of C. californica 
and A. lajollaensis on large subtidal reefs suggests some sort of negative interaction. 
Pequegnat (1964) found that C. californica was most abundant near the top of a 
subtidal reef in southern California, and became more sparse with depth. In contrast, 
A. lajollaensis formed large colonies near the base of the reef and decreased in abun- 
dance with height, occurring at low densities near the reef top. These inverse patterns 
of abundance also can be observed on large (2-5 m high) subtidal reefs at HMLR 
in central California (pers. obs.). Efforts are currently underway to document the 
distributions of these anthozoans at HMLR, and to test the ecological effects of their 
behavioral interactions in the field. 

Members of the genus Corynactis produce clonal aggregations in tropical and 
temperate marine habitats throughout the world (Carlgren, 1949). Corynactis viridis 
is the most abundant sessile organism on shallow subtidal rocks walls at the Glenan 
Archipelago on the Atlantic coast of France (Castric-Fey et ai, 1978), and at Plym- 
outh, England (Forster, 1958); C. parvula occurs on Caribbean reefs where it may 
interact with a variety of corals and sea anemones (den Hartog, 1980). These conge- 
ners may show aggressive behavior similar to that of C. californica, as well as use 
other competitive mechanisms (den Hartog, 1 977), thus affecting the abundance and 
distribution of co-occuring sessile organisms. 

Many so-called lower animals exhibit complex aggressive behaviors associated 
with resource defense. Such behaviors are observed in polychaete worms (Evans, 
1973; Dimock, 1974; Roe, 1975), chitons (Chelazzi et ai, 1983), limpets (Stimson, 
1970; Branch, 1975; Wright, 1982), sea urchins (Schroeter, 1978; Maier and Roe, 
1983), and sea stars (Menge and Menge, 1974; Wobber, 1975), as well as in the many 
anthozoans discussed in this paper. Yet the behaviors of these organisms are rarely 
considered in theoretical works on aggression and territoriality, most of which focus 
on birds and mammals ( Waser and Wiley, 1 979; Murray, 1981; Davies and Houston, 
1984), nor are they included in recent texts on animal behavior (Huntingford, 1984; 
Ridley, 1986). Because marine invertebrates often are sessile or slow-moving, and 
may be clonal as well, their aggressive behaviors have developed under a different set 
of constraints than have those of most vertebrates. More extensive consideration of 
aggression in the lower invertebrates may lead to important new insights into the 
evolution and ecology of animal conflict. 

ACKNOWLEDGMENTS 

I thank C. Hand, R. Caldwell, W. Sousa, J. Pearse, D. Fautin, B. Rinkevich, A. 
Johnson, and T. Hunter for constructive criticism throughout this research and for 



CORALLIMORPHARIAN BEHAVIOR 123 

comments on the manuscript. I am also indebted to members of Cordell Bank Expe- 
ditions, especially R. Schmieder, who collected the specimens from Cordell Bank. 
Research facilities were provided by Bodega Marine Laboratory, Joseph M. Long 
Marine Laboratory, and Hopkins Marine Station. Financial support was generously 
provided by the Lerner-Gray Fund of the American Museum of Natural History, the 
Chancellor's Discretionary Fund and a Regent's Fellowship from the University of 
California at Berkeley, Sigma Xi, and the intercampus program of the Institute of 
Marine Resources at Scripps Institution of Oceanography. This research was com- 
pleted in partial fulfillment of the requirements for the doctoral degree at the Univer- 
sity of California, Berkeley. 

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DEVELOPMENT, METAMORPHOSIS, AND SEASONAL ABUNDANCE 
OF EMB >S AND LARVAE OF THE ANTARCTIC SEA URCHIN 

STERECHINUS NEUMAYERI 

HO BOSCH, KATHERINE A. BEAUCHAMP, M. ELIZABETH STEELE, 

AND JOHN S. PEARSE 

Institute of Marine Science, University oj California, Santa Cruz, California 95064 

ABSTRACT 

The development to metamorphosis of the shallow-water antarctic sea urchin, 
Sterechinus neumayeri, is described for the first time. Developmental stages are sim- 
ilar to those of closely related temperate species with feeding larvae, but the rate of 
development is extremely slow. Hatching of ciliated blastulae occurs approximately 
140, 128, and 1 10 hours after fertilization at -1.8, -1.0, and -0.5C, respectively, 
more than twice the time required for closely related temperate species near their 
normal ambient temperature. Larvae reared at 1 .8 to 0.9C are capable of feeding 
20 days after fertilization and are competent to metamorphose after 1 1 5 days. Early 
cleavage embryos, blastulae, gastrulae, and prism larvae of this species were collected 
from the plankton adjacent to McMurdo Station, Antarctica, in early November and 
December, 1984 and 1985. Echinoplutei were not found during this study, but they 
have been collected from the plankton in other years; there is no evidence that the 
larvae are demersal. The timing of spawning ensures that feeding larvae are in the 
plankton during the abbreviated summer peak of phytoplankton abundance in Mc- 
Murdo Sound. Recruitment of juveniles into the benthos most likely occurs in syn- 
chrony with the subsequent period of high levels of benthic chl a concentrations. 

INTRODUCTION 

The perception that brooding is the prevalent mode of development among spe- 
cies of antarctic echinoderms has been firmly established over the past century 
(Thomson, 1876; Thorson, 1950, Mileikovsky, 1971; Dell, 1972; White, 1984). 
Brooding is most apparent within the shallow-water echinoid faunas (Arnaud, 1974; 
Picken, 1980). Three families in three separate orders (Cidaridae, order Cidaroidea; 
Schizasteridae, order Spatangoida; and Echinidae, order Echinoidea) represent the 
antarctic echinoids. Two of the three families are dominated by species that brood. 
Fell (1976) reported that 12 of 19 known species of antarctic cidarids are known 
brooders, and 3 others almost certainly brood. In addition, females of all 21 known 
species of antarctic schizasterids brood their young in specialized sunken aboral petal- 
oids known as marsupia (Fell, 1976). 

It is unclear whether the high incidence of brooding species among these two 
families is a consequence of ongoing selection in the antarctic environment or of 
phylogenetic history (Dell 1972; Fell, 1976; Arnaud, 1977). Fell (1976) hypothesized 
that the ancestral forms of antarctic cidarids (ancestral goniocidarids) brooded their 
young, and he suggested that cidarids colonized the antarctic as brooders. No ances- 
tral form has been clearly established for antarctic schizasterids. Since extant non- 



Received 25 September 1986; accepted 27 May 1987. 

126 



ANTARCTIC ECHINOID REPRODUCTION 127 

antarctic representatives of this group have unprotected development, the brooding 
habit of antarctic species may have evolved subsequent to their colonization of the 
antarctic (Fell, 1976). In either case, the numerical success (i.e., number of species) 
of cidarids and schizasterids in the antarctic apparently is related at least in part 
to their brooding habits. 

In contrast to the cidarids and schizasterids, the antarctic echinids are represented 
by only five species, all within a single genus, Sterechinus (Fell, 1976). Individuals of 
one species, S. neumayeri, are the most abundant echinoids in shallow-water sur- 
rounding the antarctic continent. The relatively small maximum egg sizes reported 
for three antarctic species of Sterechinus (0.15 in S. neumayeri, and 0.25 mm S. 
agassizii and .S. antarcticus) are indicative of a free-swimming mode of development 
(Mortensen, 1909, 1910; Pearse and Giese, 1966). Moreover, despite frequent collec- 
tions, brooding has not been reported for any of the six species of the genus (Fell, 
1976). The absence of post-spawning parental care among antarctic representatives 
of this group is in sharp contrast with the predominant mode of development in other 
antarctic echinoids. 

Little is known about the embryonic and larval stages of non-brooding antarctic 
echinoids. Mortensen (1913) described echinoplutei from plankton samples collected 
by the German South Polar Expedition ( 1 90 1 - 1 903). Mortensen assigned the larvae 
to 5". neumayeri because it was a common species, had very small eggs, and was not 
known to brood. A pair of echinoplutei was collected from midwater in McMurdo 
Sound by the British National Antarctic Expedition (MacBride and Simpson, 1908). 
Mortensen (1913) also assigned these to S. neumayeri. Since the publication of these 
reports over 70 years ago, little additional information on the developmental stages 
of non-brooding antarctic echinoids has been obtained. Pearse and Giese (1966) de- 
scribed the reproductive cycle of a population of S. neumayeri in McMurdo Sound, 
and suggested that the larvae of this species are demersal and not pelagic because they 
have been taken from the plankton so rarely; however, the larval development of 
antarctic echinoids had not been observed or described. 

The present paper describes the development through metamorphosis of Sterechi- 
nus neumayeri, and draws special attention to the slow rates of embryonic and larval 
development. In addition, we present information on the seasonal abundance of em- 
bryos and larvae of this species in the near-shore waters of McMurdo Sound, Ant- 
arctica. 



MATERIALS AND METHODS 

Individuals of Sterechinus neumayeri were collected by scuba divers from 15-25 
m depth beneath the annual sea ice adjacent to McMurdo Station, Antarctica (77 
51' S, 166 40' E). In November 1983, immediately after collection, approximately 
two dozen animals were transported to the University of California, Santa Cruz, 
where gametes were fertilized and the larvae were reared through metamorphosis in 
an ice bath (-0.5 to 0.5C) kept in a 4C refrigerated unit. Additional studies of the 
developmental stages and developmental rate of S. neumayeri were carried out at 
McMurdo Station; ripe animals were collected in November, 1984, and larvae were 
reared through metamorphosis and early juvenile stages to December, 1985. The 
running seawater system at McMurdo Station maintains aquarium seawater temper- 
atures between 1.8 (winter) and 0.9 (summer)C, which allowed us to rear em- 
bryos and larvae close to their ambient temperatures. 



128 I- BOSCH ET AL. 

Spawning ami >n of gametes 

Spavv .'ed by intracoelomic injection of 0.5 M KC1 solution. Eggs 

collected f spawning females were washed in clean 5 ^m filtered seawater 

an <j a few drops of dilute sperm suspension in a 4 liter polycarbonate 

ter approximately 20 minutes, eggs were filtered off with 20 /urn nitex 

ncl placed in 4 liter culture vessels with clean 5 /urn filtered seawater. 

Rearing of embryos and larvae 

Embryos and larvae were reared in gently stirred and unstirred cultures (Hine- 
gardner, 1 969; Strathmann, 1971). The water in the culture vessels was changed every 
four days using a 20 ^m mesh nitex strainer to retain the embryos and larvae. 

At McMurdo Station, larvae were fed semi-daily with equal amounts of bacterized 
cultures of Isochrysis galbana and Phaeodactylum tricornutum (total concentration 
10,000-15,000 cells/ml), which were grown at 15C in continuous light using half 
strength F medium (Guillard and Ryther, 1962). Algal samples to be used as food 
were centrifuged for 10 minutes at 5000 rpm and resuspended in clean filtered seawa- 
ter (- 1 .5C). Phytoplankton concentrations were measured using a Palmer Maloney 
counting chamber. 

Initially, at Santa Cruz, several phytoplankton species (including both temperate 
and antarctic forms) were tested as potential sources of food for the larvae. Among 
five temperate species tested (Amphidinium carter! , Dunaliella tertiolecta, Isochrysis 
galbana, Phaeodactylum tricornutum and Rhodomonas sp.), I. galbana and P. tricor- 
nutum were most resistant to low temperatures. These phytoplankton appeared to be 
healthy, even after being in larval cultures for two days, and were readily consumed 
by the larvae. Cells of the antarctic diatom Thalassiosira antarctica were not readily 
ingested by early stage plutei. 

Settlement and metamorphosis of larvae reared in Santa Cruz was induced by 
adding echinoplutei to glass dishes containing pieces of PVC pipe covered with a 
bacterial-algal film (Hinegardner and Tuzzi, 1971). The bacterial-algal film was pre- 
pared by placing the PVC pipe in a large dish that was held in a running seawater 
table for several days. Competent echinoplutei reared at McMurdo Station were suc- 
cessfully induced to settle and metamorphose with sediment samples collected from 
various depths ( 1 2, 20, 25, and 30 m) within the adult habitat. 

Embryonic developmental rates 

Time of development to hatching at different temperatures was determined for 
embryos reared at McMurdo Station by holding them in culture vials that were ( 1 ) 
in a refrigerated unit at -0.5 (-0.7 to -0.3)C, (2) in a running seawater table in the 
laboratory at - 1 .0 (- 1 .2 to -0.9)C, and (3) submerged in the sea 5 m below the level 
of the sea ice at ambient temperature, - 1 .8 (- 1 .9 to - 1 .7)C (a small heated hut with 
a hole in the floor and through the sea ice was used as a staging area). Approximately 
50 newly spawned eggs from a pair of females and a single drop of dilute sperm 
suspension were mixed in each of 3 sets of 10, 5 ml capacity vials filled with 5 ^m 
filtered seawater at the appropriate temperature. Progress of development and incu- 
bation temperature were monitored every 12-16 hours during early cleavage stages 
and every 2 hours near the time of hatching. Because agitation and small changes in 
temperature may adversely affect rates of embryonic development, only previously 
undisturbed culture vials were used for observations of developing embryos. The time 



ANTARCTIC ECHINOID REPRODUCTION 129 

of hatching was defined as the time when at least 10% of the ciliated blastulae in a 
particular incubation vial were released from the fertilization membrane. 

Field collection of embryos and larvae 

Plankton samples were collected on a weekly or bimonthly basis from September, 
1 984 to December, 1 985 using both diver-towed and stationary current-fed plankton 
nets (240 ^m mesh) at various locations in McMurdo Sound. The conical, stationary 
nets measured 2 m in length with a circular mouth opening of 0.3 m. The diver-towed 
net was 2 m long and had a rectangular mouth of 0.1 X 0.3 m. Each of the current- 
fed nets was held open continually by a steel frame; net bouyancy was regulated with 
a float. Two or three nets were attached to a weighted steel cable and suspended by 
scuba divers from the undersurface of the sea ice for 24 to 48 hours. At the points of 
attachment to the cable, the nets had a ball bearing swivel which allowed them to 
orient to the shifting directions of the prevailing currents. 

Because the larvae may be demersal, 5 replicate bottom cores of 8 cm diameter 
were taken monthly from October, 1984 through October, 1985 at 10, 20, 25, and 
30 m depth adjacent to McMurdo Station. 

All samples were sorted for larvae and other organisms within two days of collec- 
tion. Early developmental stages that were not readily identifiable were isolated from 
field samples and reared in the laboratory until they reached a recognizable larval 
stage. Sizes of embryonic and larval stages as well as larval skeletal morphology of 
the field-collected specimens were noted and compared to those of embryos and lar- 
vae reared in the laboratory from fertilization. 

RESULTS 
Sequence oj development 

Development of Sterech inns neumayeri was followed through metamorphosis at 
Santa Cruz (-0.5 to 0.5C) and McMurdo Station (-1.8 to -0.9C) (Table I). The 
eggs are small (mean diameter = 0.179 mm; n = 55) and negatively buoyant. Early 
development yields a typical sea urchin prism larva. Stomadeal breakthrough occurs 
20 days after fertilization at approximately 1.5C, and soon thereafter the larvae 
begin to feed. By the 2 1st day, the postoral and anterolateral paired arms of the echi- 
nopluteus are formed. The larval epithelium is now sparsely covered with red pig- 
ment granules, more or less randomly distributed. Formation of the posterodorsal 
and the much shorter preoral pair of arms begins at approximately 43 and 56 days 
after fertilization, respectively. The onset of the eight-arm pluteus stage is closely 
timed with the formation of the anterior epaulettes as well as the appearance of the 
five lobes of the hydrocoel (Fig. 1 ). At this stage of development the larvae are similar 
to those previously described from collections of earlier antarctic expeditions (Mac- 
Bride and Simpson, 1908; Mortensen, 1913). 

Further thickening of the ciliary band along the posterior margin of the larva 
results in the formation of the posterior epaulettes. By approximately the 80th day of 
development at - 1 .8 to 0.9C, the tube feet primordia are formed. Soon thereafter, 
a variable number (1-3) of triradiate spines appear on the external surface of approxi- 
mately 40% of the larvae. The most conspicuous of the spines is located in a medial 
position at the posterior end of the larva, while the other two are formed on the right 
side, near the bases of the postoral and posterodorsal rods of the larval skeleton. 

Metamorphosis is relatively slow, lasting 2-3 hours before the non-feeding ben- 
thic juvenile is formed. Newly metamorphosed juveniles retain many of the pigment 



1 30 I. BOSCH ET AL. 

TABLE I 

Develop^. .nm.ximate sizes of developmental stages of Sterechinus neumayeri reared 

in Santa > (-0.5 to 0.5 C) and McMurdo Station, A ntarctica (-1.8 to- 0. 9C) 







First appearance 


(days) 




Size 






)pmerital stage 


(mm) 


-1.8to-0.9C 


-0.5to0.5C 


, /edegg 


.18-.19 








blastula 


.21 


2.1 


1.7 


Hatching 





5.1 


3.7 


Gastrula 


.22 


10 


8 


Prism 


.32 


16 


15 


Early pluteus 


.35 


21 


17 


Six-arm pluteus 


.54 


43 


29 


Early eight-arm pluteus 


.80 


56 


42 


Late eight-arm pluteus 


1.20 


103 


100 


Juvenile 


.44 


115 


107 



Sizes represent the diameter of ova, blastulae and juveniles, maximum length of gastrulae and prism 
larvae, and length from the aboral apex to the tips of postoral arms of echinoplutei. 



granules characteristic of larval stages, but otherwise have a pale, whitish appearance. 
They have a single set of well developed tube feet as well as 1 juvenile and 1 5 primary 
spines. The triradiate spines which appeared on the surface of echinoplutei are re- 
tained on the aboral surface of juveniles. 

Duration of embryonic development 

Embryos reared below the sea ice (-1.9 to -1.7C), in a seawater table (-1.2 to 
-0.9)C, and in a refrigerator (-0.7 to -0.3C) at McMurdo hatched at 140, 122, and 
1 10 hours, respectively. Time to first hatching for embryos reared in an ice bath at 
Santa Cruz (0.5 to 0.5C) was approximately 88 hours. 

Occurrence of eggs, embryos and larvae in the plankton 

One hundred and twenty (120) plankton samples were taken from McMurdo 
Sound between September, 1984 and December, 1985. Of these, 56 were taken from 
near the undersurface of the ice or, in the absence of sea ice, near the surface of the 
water. Fourteen were taken from midwater (10-20 m depth), and 50 were collected 
from near the bottom at 1 5-30 m depth. 

Large numbers (500-600) of embryos, free-swimming blastulae, and gastrulae 
that closely resembled those of laboratory reared Sterechinus neumayeri were col- 
lected from the plankton at all depths sampled using both stationary and diver-held 
plankton nets. Eggs and early stage embryos were collected predominantly during the 
third and fourth weeks of November, 1 984 and 1 985. Hatched blastulae and gastrulae 
at various stages of development were predominant during the first week of Decem- 
ber, although several un hatched and newly hatched blastulae were collected from 
surface waters on the 9th of November, 1985. Four prism larvae were identified from 
midwater samples taken in mid to late December, but no echinoplutei were collected 
during this study. No sea urchin eggs, embryos, or larvae were found in the 240 bot- 
tom cores collected and examined. 



ANTARCTIC ECHINOID REPRODUCTION 



131 




100 (jm 



FIGURE 1 . Early eight-arm pluteus of Sterechinus neumayeri shortly after the formation of the pre- 
oral pair of arms (indicated by arrow). Scale bar = 100 



DISCUSSION 
Embryonic and larval development 

Compared to other species that have been studied, the developmental stages of 
Sterechinus neumayeri are most similar in shape and size to those of the temperate 
echinoid, Echinus esculentus (MacBride, 1903). However, the formation of spines 
on the external surface of the larvae, separate from the juvenile rudiment, clearly 
distinguishes the larvae ofS. neumayeri from those of E. esculentus and other species 
studied within the family Echinidae (MacBride, 1903; Arrau, 1958; Cram, 1971). 
Morphologically similar spines reportedly develop on the echinoplutei of several 
other species of regular echinoids, including both euechinoid and cidaroid forms 
(Onoda, 1931, 1936; Fukushi, 1960; R. Emlet, pers. comm.). 

The time of development for the entire period from fertilization to metamorpho- 
sis of Sterechinus neumayeri is extremely long. Within the family Echinidae, the tern- 



132 



I. BOSCH ET AL. 






UJ 

S 









90- 



60- 



30- 




10 



1 5 



20 



25 



30 



TEMPERATURE (C) 

FIGURE 2. Duration of embryonic development to hatching as a function of temperature for seven 
species of echinids and strongylocentrotids with indirect development. Hatching occurs at the ciliated 
blastula stage. Mean diameter of ova ranges between 80 (Strongylocentrotus purpuratus) to 179 ^m (Stere- 
chinus neumaveri). 4 Strongylocentrotus droebachiensis reared at 0, 4, 8C (Stephens, 1980) and 9-10C 
(Strathmann, 1974); S.frantiscanw reared at 10, and 12-13C (Strathmann, 1974); A S. pulcherrimus 
reared at 25-27C (Onoda, 1 936); O S. purpuratus reared at 1 0C (Strathmann, 1 974); D Loxechinus a/bus 
reared at 13-14C (Arrau, 1955); Parechinus angitlosus reared at 15C (Cram, 1971); Sterechinus 
neumaveri reared at - 1 .9 to - 1 .7, - 1 .2 to -0.9, -0.7 to -0.3, and -0.5 to 0.5C (this study). 



perate species Parechinus angulosus and Psammechinus miliaris are competent to 
metamorphose 60 days after fertilization at ambient temperatures (10-1 6C) (Shearer 
el al, 1913; Cram, 1971), less than half the time required for S. neumaveri near 
their normal ambient temperature (- 1 .5C). This observation agrees with the general 
trend noted by Emlet el al. (in press) between decreased temperatures (and increased 
latitudes) and increased time to metamorphosis for echinoids with planktotrophic 
larvae. 

Because factors unrelated to temperature may influence rates of post-embryonic 
development (e.g., larval food and density, Kume and Dan, 1968; Hinegardner, 
1969), we critically compared the rates of embryonic development to the hatched 
blastula stage at different temperatures, both of Sterechinus neumaveri and other sea 
urchin species with planktotrophic larvae within the families Echinidae and Strongy- 
locentrotidae. Time to hatching ranged from a minimum of 13 hours at 25-27C in 
the tropical species Strongylocentrotus pulcherrimus to a maximum of 140 hours at 
-1.9 to 1.7C for S. neumaveri, and was intermediate for temperate species near 
their normal ambient temperatures. The duration of embryonic development to 
hatching for these seven echinoid species is a curvilinear function of temperature, 
with increased sensitivity at lower temperatures (Fig. 2). A direct relationship be- 
tween the duration of embryonic development and temperature has been found with 
interspecific comparisons among other poikilotherm groups, including asteroids 
(Pearse, 1969), amphipods (Bregazzi, 1972), barnacles (Patel and Crisp, 1960), cope- 
pods (McClaren el al., 1969), and rotifers (Herzig, 1983b). Moreover, studies on sin- 
gle species or physiological races reveal the same function, describing the immediate 
thermodynamic effect of temperature on developmental processes [See for example. 



ANTARCTIC ECHINOID REPRODUCTION 133 

Bougis (1971), Stephens ( 1 972), and McEdward (1985) for temperate echinoids; Her- 
zig(1983a) forcopepods; Herzig(1983b) for rotifers; and Ross and Quetin (1986) for 
antarctic krill]. The direct relationship between temperature and duration of embry- 
onic development, both within a single species and among groups of related species, 
suggests that there is little or no temperature compensation for developmental rates 
in poikilotherms, resulting in the observed general trend of increasingly longer peri- 
ods of development with greater latitude. 

The tendency for increased lecithotrophic development among high latitude ma- 
rine invertebrates was well documented by Thorson (1950) who proposed that the 
combination of low temperatures which act to increase development time and a 
short season of phytoplankton abundance in high latitude environments select 
against planktotrophic larvae. Thorson's (1950) explanation has been challenged by 
several authors. In particular. Underwood ( 1 974) and Clarke ( 1 982, 1 983) argue that 
there should be no a priori reason to expect ontological processes to be rate-limited 
by temperature because all poikilotherms have evolved the capability to modify those 
processes for the effects of temperature. However, although numerous mechanisms 
for metabolic temperature compensation have been identified (Hochachka and Som- 
ero, 1984), there are few examples of developmental rate compensation for tempera- 
ture in any previous work (Clarke, 1 982). Development is a complex, highly synchro- 
nized process involving many biochemical and structural changes. As suggested by 
Patel and Crisp ( 1 960), basic patterns of temperature-developmental rate interactions 
may not be readily modified in evolution. 

Seasonal abundance and distribution 

The presence of embryonic and early larval stages of Sterechinus neumayeri in 
the plankton during early to mid November and December, 1984 and 1985 is in 
accordance with previous estimates of the spawning time of this species in McMurdo 
Sound (Pearse and Giese, 1 966). Observations of spawning urchins further substanti- 
ate this conclusion: males spawned in shallow water near McMurdo Station on two 
occasions during the first week of November, 1984 (B. Gullikson and T. Klinger, 
pers. comm.). Coupled with known development times of laboratory-reared embryos 
and larvae, this evidence suggests that larvae of S. neumayeri feed between late De- 
cember and early March, coinciding with the summer peak of phytoplankton abun- 
dance in McMurdo Sound (Bunt, 1964; Rivkin et a/., 1986). Consequently, settle- 
ment of larvae onto the benthos will occur predominantly during late February and 
March, in synchrony with the annual period of high benthic chl a concentration that 
occurs during the austral Fall (Berkman et al. 1986). 

Twenty-five plankton tows and 16 bottom cores were collected and examined 
between late December and early March, 1 984- 1 985, yet no echinoplutei of Sterechi- 
nus neumayeri were found. Littlepage (1966, 1968, and pers. comm.) collected and 
analyzed 547 plankton samples taken throughout the year from McMurdo Sound 
but found no echinoderm larvae. The conspicuous absence of echinoplutei from 
plankton samples taken over areas where adult 5". neumayeri are abundant led Pearse 
and Giese ( 1 966) to suggest that the embryos and larvae of this sea urchin are demer- 
sal. However, large numbers of S. neumayeri embryos and early larvae were collected 
from the water column during this study. Moreover, echinoplutei of this species have 
been taken from the antarctic plankton in other years: all 48 specimens recorded by 
MacBride and Simpson (1908) and Mortensen (1913) were taken from the water 
column; in addition, four echinoplutei of S. neumayeri were collected from near-sur- 
face waters, over approximately 300 m of water, in early January, 1986 (Rivkin et 



1 34 I- BOSCH ET AL. 

al, 1986). Thi otrates that embryos and larvae of S. neumayeri are 
readily car bottom by currents. Given the active swimming behavior 
of echi; ory cultures (I. Bosch, pers. obs.), it is unlikely that develop- 
ment of th is demersal. More extensive, multi-annual sampling is needed to 
provide ; e evidence on the larval distribution of S. neumayeri. 

ACKNOWLEDGMENTS 

We thank R. L. Britton and B. Marinovic for assistance in the field, J. S. Oliver 
for collecting and transporting urchins to Santa Cruz, and G. Fryxell for providing 
stock cultures of antarctic phytoplankton; the Antarctic Services Inc., of ITT, espe- 
cially J. Wood and S. Ackely, the Antarctic Support Services of the National Science 
Foundation, and the U. S. Navy Antarctic Support Force for their logistic support; 
W. T. Doyle, Director of the Institute of Marine Sciences, University of California, 
Santa Cruz and R. T. Hinegardner for encouragement and support; and E. Bay- 
Schmith, R. B. Emlet, R. T. Hinegardner, J. B. McClintock and J. Ott for suggestions 
on the manuscript. Supported by NSF Grant No. DPP-83 17082. 

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Reference: Biol. Bull. 173: 136-159. (August, 1987) 



FEEDING >l: \TIONS OF THE FORAMINIFERAN CIBICIDES 
REF; NG EPIZOICALLY AND PARASITICALLY ON THE 

ARCTIC SCALLOP ADAMUSSIUM COLBECKI 

STEPHEN P. ALEXANDER AND TED E. DELACA* 

A-002, Marine Biology Research Division, Scripps Institution of Oceanography, 

La Jolla, California 92093 

ABSTRACT 

The calcareous foraminifer Cibicides refulgens is a conspicuous and abundant 
component of the epifaunal community living on the valves of the free-swimming 
Antarctic scallop, Adamussium colbecki. Examination of this association using light 
microscopy, scanning electron microscopy, radiotracer, and resin-casting/sectioning 
techniques, demonstrates that the foraminifer possesses a combination of morpho- 
logical and physiological adaptations, unique among benthic calcareous foramini- 
fera, which enhance its ability to acquire nutrients in an otherwise oligotrophic and 
seasonal environment. Three distinct modes of nutrition are employed: (1) grazing 
the algae and bacteria living upon the scallop shell surface, (2) suspension feeding 
through the use of a pseudopodial net deployed from a unique superstructure of ag- 
glutinated tubes which form an extension to the calcareous test, and (3) parasitism 
by eroding through the scallop's shell, and using free amino acids from the highly 
concentrated pool in the extrapallial cavity. 

INTRODUCTION 

A variety of benthic foraminiferal species are known to live most, or part of their 
lives, epizoically on a wide range of organisms. Examples include Rosalina globularis 
d'Orbigny, R. anomala Terquem, R. carnivora Todd, Cibicides refulgens Montfort, 
C. lobatulus Walker and Jacob, C. pseudoungerians (Cushman), Discorbis wrighti, 
and Discorbinnella sp. which firmly attach to macroscopic algae and metazoans such 
as hydroids, bryozoans, tunicates, crustaceans, isopods, amphipods, decapods, pyc- 
nogonids, brachiopods, gastropods, and bivalves (Nyholm, 1961; Todd, 1965; De- 
Laca and Lipps, 1972; Hayward and Haynes, 1976; Zumwalt and DeLaca, 1980; 
Mullineaux and DeLaca, 1984; Alexander, 1985; Moore, 1985). 

The most important association in terms of numbers of foraminifera appears to 
involve filter feeding invertebrates, particularly free-swimming bivalves. Hayward 
and Haynes ( 1 976) reported 998 individual foraminifers on one specimen of the com- 
mercial scallop Clamys opercularis (Linneaus) of which 765 were Cibicides lobatulus. 
Similarly, Mullineaux and DeLaca (1984) noted an average of 1 386 foraminifers on 
21 specimens of the Antarctic scallop Adamussium colbecki, 901 of which were C. 
refulgens. 

For the majority of associations between filter feeding invertebrates and epizoic 



Received 27 March 1987; accepted 22 May 1987. 

* Presently: Division of Polar Programs, National Science Foundation, 1800 G Street, Washington, 
DC 205 50. 

136 



C. REFULGENS, MORPHOLOGY AND ECOLOGY 137 

foraminifers, the host shell may provide not only a firm substrate for attachment, but 
also the added advantage of a relatively silt-free environment. In motile bivalves liv- 
ing in areas of strong currents and wave action, the shell may provide further protec- 
tion against sand shifting and possible burial with fatal consequences (Dobson and 
Haynes, 1973; Hayward and Haynes, 1976). Even sessile molluscs such as Mytilm 
can provide a relatively silt-free substrate (Allen, 1 953), and Notocorbula living in the 
silt-laden Mississippi delta, offers a preferred habitat for Hanzawaina sp. by crawling 
above the layers of accumulating flocculent material (Bock and Moore, 1969). Fur- 
thermore, life activities of the host can enhance the availability of nutrients to the 
epizoic foraminifers; for example seven species living on the shell of the brachiopod 
Tichosina floridensis Cooper, are thought to benefit from suspended food material 
transported by the inhalant and exhalent feeding currents (Zumwalt and De- 
Laca, 1980). 

The means by which foraminifera attach themselves to the shells of their hosts 
varies considerably. Some adhere simply, with no detectable effect or marking on 
the substrate (Bock and Moore, 1969; Zumwalt and DeLaca, 1980), while others 
extensively pit and erode the subsurface layers (DeLaca and Lipps, 1972) and even 
penetrate the entire shell to reach the mantle cavity (Todd, 1965). 

In this paper we use radiotracer techniques, light and scanning electron micros- 
copy, resin casting, and sectioning methods to describe the remarkable morphological 
and physiological adaptations of a large Orbitoidacean, Cibicides refulgens, to its spe- 
cialized epizoic habit on the free-swimming Antarctic scallop, Adamussium colbecki. 

The study site 

The study site at Explorers Cove (approximately 77.6 S, 163.5 E), McMurdo 
Sound, Antarctica, was used previously by Stockton (1984) and Mullineaux and De- 
Laca (1984). 

The sediment is homogeneous fine silt mainly deposited from the late austral 
summer/early autumn freshwater input to the locality via streams originating from 
the Commonwealth and Wales glaciers (DeLaca, unpub. obs.). The virtual absence 
of currents (Mullineaux and DeLaca, 1984) prevents any reworking of the sediments 
and results in a seasonal accumulation of fine silt. Adamussium colbecki occurs in 
densities of up to 90 m 2 and forms 90% of the available hard substrate in the area 
(Stockton, 1984); it resides within depressions in the sediment which are caused by 
the light, non-locomotory flapping action of its valves (Mullineaux and DeLaca, 
1984; Stockton, 1984). 

The benthic community of Explorers Cove is thought to resemble the deep sea in 
species diversity, abundance of individual organisms, sediment relief, and long-term 
stability of temperature, salinity, and oxygen (Dayton and Oliver, 1977). The austral 
summer is accompanied by an increase in primary productivity by ice algae living 
within the lower 10 cm of sea ice, and is followed in late summer by release of the 
algae into the water column from the melting ice. Thus a seasonal pulse of organic 
material is contributed to the developing in situ productivity in shallow-water (De- 
Laca, unpub. data). 

MATERIALS AND METHODS 

Living specimens of A. colbecki were collected from 20 to 27 m by scuba diving 
through holes blasted in the 3 m thick sea ice, and maintained in an aquarium at in 
situ temperature and salinity (- 1 .8C and 34%o S). 



138 S. P. ALEXANDER AND T. E. DELACA 

Most mate >r scanning electron microscopy (SEM) was coated with gold 

and/or carbon ;>ined in a Cambridge Stereoscan MK 2 operated at 20 kV; 

images were \\ford FP4 35mm film developed in Microphen. 

Scallr-: :pifaunal communities were embedded in Spurr's low viscosity 

res j n j Inc.), then ground and polished using carborundum, diamond, 

aiK ! (,.ie abrasives until the desired plane of section was reached. Azure 

jiue, and methylene blue in borax (Richardson et al, 1960) was used 
,.:nic material. Other scallop shells, cleared of epifauna by boiling in 20 
i. n peroxide, were gradually embedded in Spurr's over three days to pene- 
trate the fine cavities of the shell material. After polymerization, the block was frac- 
tured along the plain of the shell so that the upper block retained only a thin translu- 
cent layer of calcite. Alternating treatments of 0. 1 N HC1 at room temperature, and 
3% aq. sodium hypochlorite at 60C, removed calcite and organic layers, respectively; 
the exposed face of the lower block forms a perfect cast of the scallop shell dorsal 
surface and the canals and cavities within the calcite itself. Observations were made 
with an ETEC Autoscan SEM at the Wadsworth Center for Laboratories and Re- 
search, Albany, New York. 

For SEM examination of substrate pitting, agglutinated tube morphology, and 
pseudopodial deployment, six scallops with epifauna were fixed for 2 h in 6% glutaral- 
dehyde buffered with 0. \M sodium cacodylate at pH 7.4. Dorsal valves were rinsed 
five times with distilled water, rapidly frozen at -40C, and freeze dried in an inverted 
position. Other C. refulgens were picked from all size classes of A. colbecki (Stockton, 
1984) and examined for gross morphology, aperture, and spiral face detail; attach- 
ment zones were fractured and the exposed calcite laminae studied for evidence of 
pitting or tunneling. Etched (using 0.1 N HC1) and non-etched inner valve surfaces 
were examined for perforations. 

To determine the rate of substrate pitting and agglutinated tube formation, fora- 
minifera were picked from the dorsal valves of A. colbecki, cleaned of extrathalamous 
material, and placed in semi-enclosed, transparent plastic chambers attached to areas 
of non-pitted dorsal valves from recently killed scallops (50% of which had most of 
their microflora removed). It was not possible to use living A colbecki since the frantic 
flapping of collected specimens prevented the introduction and subsequent attach- 
ment of C. refulgens to the upper valves. Such violent movements are not usual in 
the normal habit of A. colbecki (Mullineaux and DeLaca, 1984; Stockton, 1984). 
The valves with experimental foraminifers were returned to the collection site for 3 
months; the containing chambers did not alter ambient light levels and a loose fitting 
lid prevented silt accumulation but permitted exchange of dissolved materials. 

ATP analysis (DeLaca, 1986) was used to distinguish living from dead foramini- 
fera and to measure foraminiferal biomass. A carbon to ATP ratio of 300 was tenta- 
tively assumed for application to ATP values from C. refulgens, since recent work 
shows that this ratio is remarkably constant between the two taxonomically distant 
rhizopod species, Gromia oviformis (order Testacida) and Astrammina rara (order 
Foraminiferida) (DeLaca, 1 986). Cellular nucleotides were extracted with phosphate/ 
citrate buffer at 100C, and data were used for normalizing experimental results. 

Labeled amino acids ( 14 C) in the same proportions as a typical algal protein hydro- 
lysate (Amersham corporation, product CFB.25; see table I), were used to demon- 
strate uptake through the pecten shell by individual foraminifers. Plastic containers 
(12 ml volume) were sealed to the inner valve surface with silicon vacuum grease, 
and the seawater within enriched with 2.5 nCi radio-labeled amino acids at 100 nM 
final concentration. This concentration of amino acids is approximately 25 times 
lower than that recorded for free amino acids within the extrapallial cavity (see Re- 



C. REFULGENS, MORPHOLOGY AND ECOLOGY 139 

suits) and 14.7 times greater than at the sediment/water interface (DeLaca, 1982). 
Incubations lasted up to four days, and controls with containers on both faces of the 
shell (one contained label; the other retained any diffused label) measured leaching 
through non-pitted shell and passive diffusion into heat killed (30C for 30 min), 
attached foraminifers. To establish the viability of animals harvested, 10 specimens 
from each experimental group were analyzed for ATP content. 

To measure influx rates of dissolved amino acids, individual animals were re- 
moved from pecten shells, cleared of all extraneous materials, and allowed to recover 
from handling for 24 hours prior to experimentation. These animals were incubated 
in experimental medium [10 ml filter sterilized seawater (FSSW) with labeled and 
unlabeled compounds (depending upon experiment)]. Incubated specimens were 
washed in 5-6 serial baths of FSSW (~ 1 min each) until wash water registered no 
significant radioactivity over background levels. Influx was determined by measuring 
the level of accumulated radioactivity in experimental animals (homogenized in 
Aquasol 2) with a Beckman LS 6800 liquid scintillation counter. "Time zero" and 
heat killed controls were used. 

To measure grazing rates of C. refulgens, epiflora of the dorsal valves were labeled 
in situ with [ I4 C] sodium bicarbonate in light at temperatures between 1.8 and 0C 
for 1 2 hours; individual cleaned and heat killed C. refulgens were placed on these 
shells prior to labelling. After incubation, the scallop shells with foraminifers were 
washed in serial baths of FSSW until the radioactivity of wash water was not signifi- 
cantly over background levels. Twenty individual diatoms, living foraminifers, and 
heat killed foraminifers were selected as time-zero samples and extracted in 1 .0 ml of 
hot ( 100C) phosphate/citrate buffer. After removal of 10 ^1 of supernatant for ATP 
analyses, the extract was dried and digested with 0.3 ml of 0.2 TV perchloric acid prior 
to adding Aquasol 2. Subsequent specimens were sampled at three 6-h intervals, and 
similarly processed. Radiation counts obtained from the time zero specimens were 
subtracted, and the results used to calculate the number of cells ingested. Experimen- 
tal protocols for isolating, washing, and determining 14 C uptake by single algal cells 
were taken after Rivkin and Seliger (1981). 

Suspension feeding was investigated using radiolabeled bacteria and diatoms. 
Bacteria were isolated from the sediments of New Harbor and further isolated on 
2216 Marine Agar (Difco). Selected cultures were then labeled with [ I4 C] leucine 
(ICN) at log growth in Marine Broth (2216 Difco). Labeled bacteria were washed 
free of extraneous label by repeated centrifugation and resuspension in FSSW, until 
supernatant radioactivity was not significantly over background levels. Cell concen- 
trations were determined with a Petroff-Hauser counting cell, and disintegrations per 
cell measured by liquid scintillation using Aquasol 2. Nitzchia cylindricus cultures, 
provided by Dr. C. W. Sullivan (University of Southern California), were grown with 
Alga-grow media (Carolina Biol. Suppl. Co.) in FSSW and labeled with [ I4 C] sodium 
bicarbonate. These cultures were concentrated and washed on nitex screen and resus- 
pended to the desired concentration (measured using a plankton counting cell). 

Four dorsal valves of living Adamussium colbecki were removed, and the aggluti- 
nated portions of 50 C. refulgens were gently but thoroughly removed from the test, 
leaving the foraminifers securely attached to the shells. Two of the shells were main- 
tained at temperatures between - 1 .8 and 0C, and the others were warmed to 30C 
for 30 min (heat killed controls); shells were suspended upside down for 6 h in a 
culture vessel with a suspension (maintained with a small stream of air) of labeled 
diatoms in seawater. This configuration was duplicated to measure bacterial capture. 
At to and hourly intervals, samples of both diatom and bacterial suspensions were 
taken (by centrifugation and filtration with nitex screen, respectively) to check for 



140 P ALEXANDER AND T. E. DeLACA 

dissolved label a asure cell concentration. Following incubation, 30 foramin- 

ifers with <, -"s, and 30 foraminifers without, were detached, cleaned 

of extranet rial i. if necessary), washed through 10 serial washings of FSSW, 

extractec phosphate buffer, and processed as described above. 

were measured using light microscopy. Approximate cell vol- 
umes -'<ed by appropriate geometric formulae corresponding to the cell 

content was also calculated (Strathmann, 1967). 

/olumes of fluid were sampled from the extrapallial cavity of A. colbecki 
een the mantle and inner surface of the shell) by passing a blunt cannula at- 
iied to a syringe through a window cut in the 0.5-0.7 mm thick shell. Separation, 
identification, and quantification of free amino acids in selected samples were accom- 
plished using high pressure liquid chromatography (HPLC) after fluorescence derivi- 
tization with ortho-pthaldialdehyde (see Stephens, 1982). 

RESULTS 

The dorsal surfaces ofAdamussium colbecki living in sedimentary environments 
are encrusted with attached foraminifera (Fig. 1) including C. refulgens. The force 
required to dislodge an individual of C. refulgens from the surface of a bivalve shell 
increases with size of the individual, as does the extent and depth of substrate erosion. 
Juvenile C. refulgens (Fig. 2) are easily dislodged using a fine needle, whereas adult 
specimens (Figs. 3-6) must be pried off with a stout microprobe. A random sample 
of shell surfaces under Cibicides refulgens demonstrated that only 45% (n = 200 on 
each of 5 shells) of foraminifers had caused etching. Similar sampling near the umbo 
revealed that 92% of the foraminifers resided in etched concavities whereas only 12% 
of those foraminifers nearer the shell margins were attached over etched shell (100 
foraminifers examined on each of 5 shells). No significant etching was detected after 
three months on shells artificially infested with C. refulgens (Fig. 7). 

A progression in the extent of substrate pitting caused by increasing sizes of fora- 
minifers and the age of the shell is clearly visible using a dissecting microscope. Use 
of the SEM demonstrated that pits caused by younger C. refulgens generally extend 
no deeper than the uppermost laminae of the valve. At this stage the striations visible 
on the shell surface (Stockton, 1984) and the smaller perpendicular 'ribs' connecting 
them (Fig. 8) are removed completely in the area of the pit, and the exposed calcite 
is eroded to appear as irregular granules with multitudinous 'micro-canals' (Figs. 9, 
10). Further pitting results in enlargement of the microcanals to form distinct canal 
openings which penetrate several calcite layers, and are to some extent guided by 
planes of weakness within, or between, the layers (Figs. 11, 13). This phenomenon is 
dramatically illustrated by resin casts which show the pit erosion in reverse, produc- 
ing a 'cathedral effect' from the pattern of channels within the scallop shell material 
(Fig. 12). The distributions of canal openings within the substrate pits as a whole do 
not exhibit any noticeable pattern. 

The bond between the test wall of the umbilical face of C. refulgens and the upper- 
most calcite layer in the pit is sufficiently strong so that when a specimen is forcefully 
detached from the shell, a layer of shell material will often remain attached to the 
foraminifer. It is then possible to observe deeper canals ramifying through the shell; 
these canals are generally fewer in number than the more superficial canals, but are 
larger in diameter ( 10-14 ^m) and more conspicuous. 

Fracturing a scallop shell directly through a substrate pit allows for detailed SEM 
study of groups of canals in the middle and lower layers (Fig. 22). Scanning electron 
micrographs (Figs. 13, 14, 16) demonstrate conclusively that the canals do not funda- 



C. REFULGENS, MORPHOLOGY AND ECOLOGY 



14 




*" 



FIGURE la. The free swimming Antarctic scallop Adamussium colbccki with characteristic epizoic 
growth. Conspicuous attached faunal components include the agglutinated tube of a large polycheate, 
hydrozoans, bryozoans, and commonly four or more species of benthic foraminifera. The most abundant 
and conspicuous species is Cibicides refulgens with its agglutinated tubes. Scale bar = 1 cm. b. Oblique 
view of the dorsal valve of A. colbecki with attached C. refulgens and associated agglutinated tubes reaching 
into the overlying water. Vertical tubes may extend to 5 mm and exhibit three orders of branching. Scale 
bar = 5 mm. 



mentally follow lines of weakness within the shell, and therefore it appears that the 
foraminifer's cytoplasm can control both the extent and direction of the dissolution 
process. The thick resin cross-sections of C. refulgens attached to the valve surface 
revealed many visible canals extending from the base of the pit through most of the 
calcite layers perpendicular to the plane of the laminae (Figs. 17, 18). However as a 



142 



S P. ALEXANDER AND T. E. DELACA 




FIGURE 2. Juvenile Cibicides refulgens attached to surface of A. colbecki dorsal valve. A rudimentary 
peripheral agglutinated tube has been built (white arrow). Vertical tubes are not present. Two juveniles 
have been removed to show the shallow surface etching of the shell (black arrows). Scale bar = 163 /mi. 

FIGURE 3. Plan view of an attached adult with a well developed agglutinated tube system (large 
arrows) and net of pseudopodia on the scallop shell surface (small arrows). Scale bar = 200 ^m. 

FIGURE 4. Oblique view of specimen in Figure 3. Pseudopodia (small arrows) can be seen traversing 
the space between the agglutinated tubes (large arrows) and the substrate. Scale bar = 143 Mm. 

FIGURE 5. Detail of Figure 3 showing composition of agglutinated tubes and the presence of a fine 
pseudopod (arrows) radiating away from the foraminifer, across the shell surface. Scale bar = 102 /nm. 

FIGURE 6. Oblique view of two attached adult Cibicides refulgens. An agglutinated tube can be seen 
clearly raised away from the scallop shell surface (arrow). Scale bar = 1 54 



C. REFULGENS. MORPHOLOGY AND ECOLOGY 143 

result of the curvature of the canals, and the limited depth of field, it was not possible 
to photograph a single element traversing the complete shell thickness without inter- 
ruption. The canal walls are significantly smoother than the adjacent calcite exposed 
at the fracture zone (Figs. 16, 22), but there is no evidence of an actively secreted 
lining. 

The inner valve surface is generally lined with overlapping, angular, tile-like cal- 
cite crystals (Fig. 23), between which are many naturally occurring pores leading to 
the lamina beneath (Fig. 24). Upon careful scrutiny of this inner layer in the SEM, 
circular areas (approximately 15-50 ^m in diameter) with significantly enlarged 
pores (Fig. 23) are evident marking the area above which an individual foraminiferan 
is attached on the outer valve surface. After etching for 5 to 10 minutes with 0.1 TV 
HC1, the outermost calcite layer is removed and those salient markings are revealed 
more clearly (Figs. 19, 20). High magnification detail shows them to be closely spaced 
canal openings (Fig. 2 1 ), and there remains little doubt that these openings are contin- 
uous with the canals which originate in the surface pit, and penetrate deep into the 
bivalve's shell. 

Examination of the exposed face of an adult C. refulgens detached from the sub- 
strate (Fig. 25, 26) reveals that the spiral face is not adhered to the shell material over 
its complete area due to a pattern of grooves radiating from the primary aperture to 
the peripheral test margins. The roof of each groove is the spiral test face, and the 
floor is formed by the etched shell material of the pit floor. Typically, four to five such 
grooves of approximately 30 to 50 nm width connect areas immediately adjacent to 
the primary aperture (Figs. 25, 29) with those more remotely situated on the opposite 
test margins. Etched bivalve shell which forms the base of the grooves often exhibits 
a pattern of fine channels (5-10 /urn wide) running parallel with the main trend of the 
groove (Fig. 30), giving the impression of the streamlines oriented with the direction 
of the main cytoplasmic flow within the grooves. In addition, the lumina of the radial 
grooves are continuous with that of the peripheral tube encircling the test at its point 
of contact with the substrate (Figs. 25, 26). 

Cibicides refulgens secondarily forms an elaborate agglutinated tube system 
around, and extending from, its attached test. The agglutinated tube system typically 
is comprised of: ( 1 ) a peripheral tube encircling most, if not all, of the lateral test 
margin at the point of contact between it and the substrate (Figs. 2, 3, 25, 26), and 
(2) radial tubes attached to the substrate and test, extending over the shell surface and 
vertically away from it (Figs. Ib, 3, 4, 27). These tubes often form several branches 
(Fig. 3). 

All radial tubes originate from the peripheral tube, either dividing from it without 
any observable thickening, or arising from a distinct node at a particular point. Tubes 
extending horizontally along the shell surface and vertically into the overlying water 
may branch from the same nodes. There is no obvious organization of the branching 
patterns of C. refulgens. Typically, C. refulgens has from 1 to 6 (x == 3, n = 50) aggluti- 
nated tubes extending vertically up to 5 millimeters (x ---- 2.5, n ---- 50) from their 
points of origin at the peripheral tube. Vertical tubes may exhibit up to three orders 
of branching, and tube diameter does not vary consistently with length or distance 
from the test/substrate; thickenings or nodes can occur at any point along a tube. The 
interior tube surface is smoother than the outer surface (Fig. 26), and in freeze dried 
specimens it is partially covered with a layer of cytoplasm. 

Intact tubes, when viewed in the SEM, do not show clearly defined apertures; 
openings are inferred by the presence of pseudopodia which extend from many points 
along the tubes to either the shell surface, test surface, or other tubes. Apparent aper- 



144 



S. P. ALEXANDER AND T. E. DeLACA 



' 



. Ti 

. . .w.i 

It; -;,%: 




FIGURE 7. Substrate markings caused by an adult Cibicides refulgens after three months of attach- 
ment. There is no visible etching of the shell surface, but adhesion was great enough to break away some 
test material upon removal of the foraminifer. Scale bar = 200 /urn. 

FIGURE 8. 'Early stages' of substrate pitting. The striae of the scallop shell have been removed, and 
from two to three laminae have been eroded. There is no evidence of boring to form canals. Scale bar 
= 110 urn. 

FIGURE 9. Detail of peripheral region of an early stage pit. The surface lamina has been etched away 
(lower left) and the calcite beneath has been partially eroded to form fine, angular granules. Scale bar = 1 1 
/urn. 



C. REFULGENS. MORPHOLOGY AND ECOLOGY 145 

tures such as that shown in Figure 3 1 are caused by tube breakage during collecting 
or transport of the scallops. 

Removal of specimens from the water causes the vertical tubes to collapse against 
and adhere to the substrate, forming what then appears to be a system of surface tubes 
which resemble polycheate worm tubes. However, cytologically fixed and freeze dried 
tubes are able to partially support themselves thereby almost maintaining their natu- 
ral positions (Figs. 4-6), and thus facilitating examination in the SEM. 

The walls of all tubes are clearly agglutinated and comprise fine (silt- and clay- 
sized) mineral particles, diatom frustules, fine organic detritus, and occasional sponge 
spicules (Figs. 5, 26-28, 3 1 ). The cementing material is not clearly distinguished from 
the agglutinated particles and does not cover their outer surfaces. Wall thickness var- 
ies considerably but is generally from one to four particles thick with no evidence of 
layering or selection of specific particle size for construction. Particle recruitment 
by foraminiferal cytoplasm during tube construction seems to be dependent on the 
availability of sedimentary material on the scallop shell surface. Similarly, the incor- 
poration of specific diatom frustules into the tubes is related to the dominant flora 
growing upon the scallop shell, and perhaps, the diet of the foraminifer. 

Extrathalamous cytoplasm and pseudopodia were studied for gross morphology 
using both living and freeze dried specimens attached to pectin shells. An extensive 
pseudopodial net was observed spread over most of the shell surface in areas densely 
populated by C. refulgem (Fig. 32). The dorsal test surface of C. refulgens usually is 
partially covered with cytoplasm in the form of tangled strands (Figs. 32, 33). From 
this, randomly branching and anastomosing networks of pseudopodia emanate and 
connect with neighboring tests, agglutinated tubes, and/or clumps of algal or detrital 
material. Trunk pseudopods are usually found closer to the substrate, originating 
from peripheral or vertical agglutinated tubes, and traversing portions of the shell 
while remaining suspended above it (Figs. 3, 4, 5). Fine pseudopodial elements 
branch at apparently random points along trunk pseudopods and connect with others 
nearby (Fig. 34 ), or attach to the substrate beneath. These elements often merge with 
a finer net system attached to the substrate at raised points such as striae, and spread 
over most of the shell surface in the vicinity of the adult C. refulgens (Fig. 35). The 
rectilinearity and patterning of elements forming the fine nets and the limited extent 
of sagging when they are bearing dense mineral particles is indicative of tension 
within the system. The larger trunk pseudopods often visibly sag when crossing spaces 
between neighboring foraminifers and clumps of detrital material. 

Vertical agglutinated tubes also give rise to trunk pseudopods and finer branching 
elements suspended freely in the water space immediately surrounding and above 
the tubes. Relative movement of the water in this space causes the pseudopods to 
bend and wave freely, demonstrating extreme flexibility in response to water 
movement. 

All of the pseudopodia have a sticky quality when touched with single hair brushes 
or steel microprobes; once adhered they stretch considerably under tension before 
breaking. Diatoms, sedimentary particles, and organic debris are commonly observed 
attached to pseudopodia (Figs. 36, 37), and large clumps of detrital material were 



FIGURE 10. Detail of pit base in Figure 9. Note the fine 'pores' visible between the angular calcite 
granules. Scale bar = 2.8 ^m. 

FIGURE 11. A well developed substrate pit (bottom half of micrograph) with characteristic deep 
borings visible (arrows) penetrating several laminae of the scallop shell. Scale bar = 62 urn. 



146 



S. P. ALEXANDER AND T. E. DnLACA 




FIGURE 12. A resin cast of the central portion of a substrate pit formed by an adult Cibicides reful- 
gens; the raised central area represents channels within the scallop shell which were originally occupied by 
foraminiferal cytoplasm. Scale bar = 20 nm. 

FIGURE 13. Irregular etching of calcite at the pit edge. A bored hole in upper calcite layer (X) has 
been undercut by subsequent dissolution of lower layers (arrow). Scale bar = 3 1 nm. 

FIGURE 14. Peripheral area of a deep substrate pit showing transition from scallop shell surface 
(bottom right) to extensively etched pit base (top left) and a circular vertical boring (X). Scale bar = 18 Mm. 

FIGURE 1 5. Transition from normal scallop shell surface (left) to a deep pit (right) eroded by a large 
adult Cibicides refulgens. Removal of the foraminifer has torn away the uppermost calcite layer, exposing 



C. REFULGENS. MORPHOLOGY AND ECOLOGY 147 

often observed suspended above the substrate within pseudopodial nets. Such inclu- 
sions may be partially engulfed by cytoplasm and/or suspended by a pattern of reticu- 
lar 'subnets' formed between main pseudopodial elements (Fig. 36). 

Figure 40 presents the results of three experiments to further examine the sources 
of particulate organic material used as a nutrient source by Cibicides refulgens. 
Though patchy in distribution, benthic diatoms (primarily Cocconeis sp. approxi- 
mately 1 5 /urn in length) represent a potentially significant resource to grazing organ- 
isms living on the surface of the bivalve. Time course studies monitoring the corn- 
sumption of radio-labeled benthic algae by C. refulgens demonstrated average grazing 
rates of 54.5 diatoms mg~' h ' (n = 60, min = 14, max = : 1030). The relatively large 
differences in uptake rate can be accounted for by the proximity of the foraminifer's 
attachment site to benthic diatoms on the surface of the bivalve shell. Two other 
experiments examined the rate of capture of suspended bacteria and diatoms. These 
experiments were additionally designed to determine the relative importance of the 
agglutinated tubes in suspension feeding. In both of these experiments half of the 
attached foraminifers were cleaned of all agglutinated tubes to evaluate the impor- 
tance of these structures to suspension feeding efficiency. While suspended cultures 
ofNitzchia cylindricus (5-10 fj.m at concentrations of 8 X 10 4 cells ml" 1 ) were taken 
at rates of 153 cells mg ' h~' (n = 15, min. = 82, max. = 284) by foraminifers with 
their arborescent agglutinated tube structures intact, those without this superstruc- 
ture averaged rates of only 62 cells mg" 1 h" 1 (n = 15, min. = 32, max. = 141). Sim- 
ilarly, suspended bacteria (unidentified gram negative rods 1.2 X 2 ^m at 2 X 10 6 
cells ml" 1 ) were consumed in greater numbers by C. refulgens with agglutinated tubes 
(x = 4.2X 10 2 cells mg" 1 h"',n == 15, min - 1.1 X 10 2 , max. = 6.7 X 10 2 ) than those 
without agglutinated tubes (x = 69 cells mg" 1 h" 1 , n = 15, min. = 21, max. = 1.7 
X 10 2 ). 

The discovery of pronounced etching channels penetrating through the shell 
clearly placed foraminiferal cytoplasm in contact with the nutrient-rich extrapallial 
space formed between the mantel and the inner surface of the shell, and suggested a 
parasitic relationship. Our studies using radio-labeled amino acids demonstrated the 
0.5-0.7 mm thick unetched bivalve shell is not permeable to free amino acids. How- 
ever, when the inner surface of the bivalve shell opposite attached C. refulgens was 
bathed with radio-labeled amino acids (100 p.M), the foraminifers consistently be- 
came radioactive within a few hours. These experiments were duplicated with heat 
(30C for 30 min) killed C. refulgens and no radioactivity was detected. 

Figure 39 presents the uptake of uniformly 14 C labeled mixture of amino acids 
from seawater at various concentrations. This curve is clearly hyperbolic and suggests 
that the transport system for amino acids in Cibicides refulgens can be described by 
the Michaelis-Menten equation. The data have therefore been analyzed by a Hanes- 
Woolf plot (where substrate concentration divided by rate of influx is plotted against 
substrate concentration). As shown in Figure 40, J max is 3.59 X 10" 3 ^moles mg" 1 
(wet weight of protoplasm) IT 1 and the K t (substrate concentration at which the rate 
of uptake = J max /2) is 10.43 \iM. 

Analysis of the fluid filling the extrapallial cavity were conducted using high pres- 



unetched laminae beneath and canals penetrating deeper into the shell material (arrows). Part of the agglu- 
tinated peripheral tube remains secured to the shell surface (large arrowheads). Scale bar = 36 nm. 

FIGURE 16. Detail of Figure 1 5 demonstrating the distinct canal borings (X) in shell material beneath 
the attached foraminifer. Scale bar = 6.7 



148 

17 



S. P. ALEXANDER AND T. E. DeLACA 




FIGURE 1 7. Thick cross section through a resin-embedded adult Cibicides refidgens attached to Ad- 
amussium colbecki. Groups of canals are discernible originating from the base of the pit and passing 
through most of the shell thickness (arrows). Scale bar = 200 ^m. 

FIGURE 18. Detail of Figure 17 showing the canals to be continuous through to the inner-most 
laminae of the scallop shell (arrows). Scale bar = 480 ^m. 

FIGURE 19. Acid etched inner shell surface with foraminifers attached to the opposing face. Each 
mark (arrows) corresponds to groups of canals penetrating the shell from the surface pits above. Scale bar 
= 250 /im. 



C. REFULGENS. MORPHOLOGY AND ECOLOGY 149 

sure liquid chromatography. The results (Table I) revealed concentrations of 2527 
nM (2.527 mM) free amino acids with extremely high concentrations of glycine 
(2066.3 /uM). 

DISCUSSION 

The rarity of loosely attached or 'roaming' C. refulgens on the surfaces of scallop 
shells strongly suggests that the sessile habit is preferred by this species. The poorly 
eroded pits beneath juveniles and the extensive pits associated with adults, leads to 
the assumption that pitting progresses with growth at least until the adult stage is 
reached (data on the life span of C refulgens are unavailable). Specimens experimen- 
tally placed on a previously unmarked scallop shell became firmly attached to the 
substrate and began to construct agglutinated tubes. However, the lack of significant 
etching after three months raises several interesting questions: is this the typical rate 
of etching which would be observed by those specimens attached to live A. colbecki? 
Alternatively, is it significantly lower in response to the absence of specific cues? We 
have demonstrated that amino acids do not normally leach through the shell mate- 
rial, thus it seems unlikely that this would act as a cue to initiate excavation; cues 
could conceivably come from a variety of stimuli, such as rates of sediment accumu- 
lation on the shell surface, the presence or absence of organic materials from scallop 
excretion, and the presence/absence of water movements over the shell surface. Ex- 
tensive further studies are required to investigate the role (if any) of environmental 
cues in initiating substrate erosion by C. refulgens. 

Parasitism 

Our investigations have shown that 50% of attached C. refulgens significantly 
erode the surface of the scallop's shell and excavate channels to the extrapallial cavity. 
Though the scallop shell normally is not permeable to dissolved amino acids, radiola- 
beled studies have consistently shown uptake of amino acids by attached foramini- 
fers. This uptake could only have been mediated by pseudopodia penetrating through 
the shell. The nutritional significance of dissolved amino acids to several marine in- 
vertebrate species has been discussed by other workers (Southward and Southward, 
1972; Stephens, 1981) including foraminifera (DeLaca el ai, 1981; DeLaca, 1982). 
Cibicides refulgens has the ability to absorb free amino acids at relatively low sub- 
strate concentrations (K, = 10.43 nM). However, concentrations of free amino acids 
within the extrapallial space are more than two hundred times higher (2527 nM, 
[2.527 mA/]) than the half saturation concentration (K t ). Therefore, a logical assump- 
tion that the foraminifer has little difficulty realizing its maximal rate of influx (J max 
= 3.59 X 10 3 Mg, mg~ 1 ,h"')from the scallop, and presumably this source of material 
would be available to the foraminifer year-round. 

A wide range of associations between individuals of different species in which one 



FIGURE 20. Detail of Figure 1 9 showing salient features directly beneath a substrate pit on the oppos- 
ing shell surface. Scale bar = 60 ^m. 

FIGURE 21. Detail of Figure 20 showing fine perforations present in laminae exposed by etching 
with HC1. Scale bar = O.9 /mi. 

FIGURE 22. Detail of canals exposed during fracture of the shell through a pit. These canals are 
approximately midway through the dorsal shell material and contain precipitated material, most probably 
cytoplasm. Scale bar = 23 



150 



S. P. ALEXANDER AND T. E. DeLACA 




FIGURE 23. Inner surface of dorsal scallop valve showing marking which is often observed when the 
opposing surface is heavily colonized with Cibicides refulgens. Scale bar = 6.2 nm. 

FIGURE 24. Detail of Figure 23 demonstrating enlarged 'pore' between calcite plates (X) and numer- 
ous 'micropores' located peripherally (arrows). Scale bar = 1 .0 ^m. 

FIGURE 25. Umbilical view of an adult Cibicides refulgens removed from the valve surface. The 
primary aperture (arrow head) opens into (a) the lumen of the peripheral agglutinated tube (dashed line) 
and (b) grooves created between the umbilical face and the base of the pit (dotted lines). Scale bar = 150 



C. REFULGENS, MORPHOLOGY AND ECOLOGY 151 

or both derive benefit from the other have been described in the literature. They range 
from being obligate to being facultative (each partner being able to live without the 
involvement of the other), and the grades of association within this range often are 
not distinct. For convenience the relationships are frequently termed commensal and 
parasitic. By definition, a parasite always lives to the detriment of its host. Parasitic 
life styles are frequently specialized and lead to development of morphological as well 
as physiological adaptations which ensure efficiency. The relationship between C. 
refulgens and A. colbecki is very similar to that described by Todd ( 1 965 ) for Rosalina 
carnivora and Lima angolensis. Unlike Todd's work however, the present study pres- 
ents unambiguous evidence that C. refulgens does derive nourishment from the man- 
tel of its host. Whether the cumulative affects of approximately 900 attached C. reful- 
gens (~400 [45%] of which may have created channels through the shell) have a 
detrimental affect on the bivalve in this marginal environment remains unknown, 
but seems likely. 

Grazing 

Morphological test elaboration in the form of a constructed horizontal tube sys- 
tems on the scallop shell surface effectively increases the distance that pseudopodia 
can gather food without severely increasing risk of cytoplasmic loss to predation or 
other causes. For example, we have observed tanaid crustaceans living in tubes on the 
scallop shell and feeding on unprotected cytoplasm of extended pseudopodia from C. 
refulgens. Of course, cytoplasm within the agglutinated tubes of the foraminifer is 
contiguous with cytoplasm in the lumen of the last formed chamber and thus the 
tubes are regarded as an extension of that chamber. Whereas most calcareous fora- 
minifera are compelled to withdraw all extrathalamous cytoplasm into the test when 
unfavorable conditions or predators are encountered, C. refulgens individuals need 
only withdraw pseudopodia into the agglutinated tubes for protection. Thus the total 
volume of cytoplasm deployed, and therefore the total area grazed, is vastly increased 
without much risk of cytoplasmic loss. Most calcareous foraminifera use only the 
cytoplasm present in the last formed, penultimate and sometimes the antepenulti- 
mate chamber for pseudopodia and extrathalamous cytoplasmic activity (Anderson 
and Be, 1978; Anderson, 1983; Alexander and Banner, 1984; Alexander, 1985), and 
therefore may gather food at limited distances from the test without considerable risk 
to cytoplasm. 

Adamussium colbecki shells typically are colonized by benthic diatoms and bacte- 
ria, but their concentrations, diversity, and percentage of surface coverage, however, 
vary from specimen to specimen. This heterogeneity is typical in New Harbor both 
spatially and temporally on large and small scales, and organic productivity in this 
portion of McMurdo Sound is extremely seasonal; our observations indicate that 
pronounced shallow-water productivity may be limited to as little as 2'/2 months. 
(DeLaca, unpub. data). Although approximately six months of continuous sunlight 



FIGURE 26. Detail of Figure 25, demonstrating continuity of the primary aperture (arrow head), 
with the lumen of the peripheral agglutinated tube (arrows) and that of a vertical agglutinated tube (V). 
Scale bar = 67 ^m. 

FIGURE 27. Surface morphology of a typical agglutinated tube (in this case, radial and in contact 
with the substrate). Scale bar = 67 /urn. 

FIGURE 28. Detail of typical agglutinated material forming tube walls. Arrows = diatom frustules. 
Scale bar = \2 



152 



S. P. ALEXANDER AND T. E. DELACA 



^ . - , 

TS " * * 

" ->- r> ' ~''V : ' 

' * . ' ^* "^ - V JW ** 

. ** -+ * ~ 




FIGURE 29. Detail of Figure 25 showing a radial umbilical 'groove' (G) which exists between the 
substrate surface in the pit and the umbilical test wall. A = primary aperture; C = Calcite broken away 
from pit base. Scale bar = 48 ^m. 

FIGURE 30. A typical eroded 'channel' (Ch) commonly observed on calcite which forms the base of 
umbilical 'grooves' in attachment pits. Scale bar = 4.2 jum. 

FIGURE 31. Detail of agglutinated tubes (T) formed by juvenile Cibicides refulgens in Figure 2. D 
= Diatom; W = test wall of last formed chamber. Scale bar = 18 ^m. 

FIGURE 32. Oblique view of adult Cibicides refulgens attached to dorsal scallop valve. An extensive 
net of pseudopodia (arrow heads) is visible over the substrate and extending from the dorsal test surface 
(arrows). Scale bar = 83 



C REFULGENS. MORPHOLOGY AND ECOLOGY 153 

is available annually, the combination of low angle of incident radiation, sea ice, and 
snow cover reduces the period of primary' productivity further (see Dayton and Oli- 
ver, 1977). 

Benthic diatoms on the scallop shell surface, as well as amorphous organic mate- 
rial and sediment, were attached to and transported by pseudopodia. Our experi- 
ments using in situ l4 C-labeled attached, and motile benthic diatoms, demonstrate 
that the foraminifers graze upon the naturally occurring 'lawns' of algae, and that 
relatively high numbers (x = 54 diatoms mg~' h~') are consumed. Through the ap- 
proximation of biomass and conversion to carbon content (Strathmann, 1967), it is 
estimated that if rate of harvest remained constant, those benthic diatoms would have 
contributed approximately 1 X 10~ 3 Mg C mg~' h '. That value is approximately one- 
half the amount of carbon obtained through the uptake of dissolved amino acids; 
thus grazing microorganisms from the surface of the scallop shell appears to be an 
important factor in C. refulgem nutrition. The discorbid foraminifer, Rosalina globu- 
laris, also forms deep pits in its preferred substrate (DeLaca and Lipps, 1972), and 
grazes upon algae in the immediate vicinity; however, in conditions of low food con- 
centration it roams in search of algae (Sliter, 1965). In view of the patchy distribution 
of algae on A. col beck i, the selection of a sedentary habit by C. refulgens seems to 
have potentially reduced its grazing abilities. This disadvantage is more than compen- 
sated for by the permanent, or semi-permanent attachment between the scallop shell 
and the foraminifer, which virtually eliminates the risk of being swept off the shell 
during swimming movements of the scallop, and enables further morphological and 
physiological adaptations to the epizoic habit. 

The radial grooves existing between the spiral side of the foraminifer' s test and the 
surface of the pits increases the efficiency of this greater cytoplasmic volume through 
provision of a short line of communication between the primary aperture and the 
lumen of the peripheral tube on the opposite test side; this facilitates rapid exchange 
of cytoplasmic organelles and inclusions such as mitochondria, phagocytosed mate- 
rial, and energy substrates between the most distal pseudopodia and the intrathala- 
mous cytoplasm. These internal-external lines of communication between deeply 
situated cytoplasm and the external milieu, are considered important in foramini- 
feran cell systems (Brasier, 1982). 

Suspension feeding 

Using the vertical agglutinated tubes as conduits for streaming pseudopodia and 
as anchors for pseudopodial nets, C. refulgens exploits suspension feeding as a third 
trophic mechanism. Figure 38 depicts the most typical arrangement of pseudopodia 
in an undisturbed living specimen; free pseudopodia are not rigid structures, but yield 
to water movement. Pseudopodial nets are randomly arranged and thus form a wide 
range of mesh sizes. Construction of the nets is initiated by the extension of pseudopo- 
dia from apertures along the vertical tubes, followed by contact with other tubes or 
nearby structures, and elaboration through bi-directional cytoplasmic flow. While 
unsupported pseudopodia of C. refulgens also have been seen projecting into the 
water, the construction of an agglutinated tube system provides scaffolding for further 
suspended pseudopodia within the water column, as well as a reservoir of protected 



FIGURE 33. Dorsal test surface (TS) of an adult Cibicides refulgens showing cytoplasmic strands 
(arrow heads) reaching to adjacent detrital material (far right). Scale bar = 3.2 nm. 



154 



. P. ALEXANDER AND T. E. DELACA 




FIGURE 34. Trunk pseudopodia (TP) crossing the scallop shell surface and radiating away from a 
large adult Cibicides refulgens. Fine pseudopodia can be seen branching from the main trunk and attaching 
to the substrate (arrows). D = diatoms. Scale bar = 16.6 /on. 

FIGURE 35. Finely branching pseudopodia (arrows) forming a net above the substrate. Detrital mate- 
rial (De) and diatoms (D) are entrained by pseudopodia. Scale bar = 1 7.2 ^m. 

FIGURE 36. Diatom (D) and attached detritus suspended above substrate by a fine anastomosing 
pseudopodia (arrows). Scale bar = 6.9 ^m. 

FIGURE 37. Cytoplasm (Cy) of pseudopod which has adhered to several diatoms (D) on the substrate 
surface. Note fine cytoplasmic threads (arrow heads). Scale bar = 8 



C REFULGENS, MORPHOLOGY AND ECOLOGY 



155 




39 



1 

J. 



3.0- 



2.0- 



1.0- 




40 



-3 



I 



K =10.43/11 M 



3 300 1 



200 



100 



10 



50 



100 



.y E 

5 2 

c ^ 

<U ^ 

CD D 



11 
.2 

5 Q 



-o 
c 



FIGURE 38. Cibicides refulgens attached to the shell of Adamussium colbecki, with pseudopodia 
deployed from agglutinated structures. (Not to scale.) 

FIGURE 39. Velocity of uptake as a function of the concentration of dissolved amino acids in seawa- 
ter. The values for J max (ngrams/mg h) and K., were obtained from a Hanes- Wolff linear transformation of 
the data. Each point is the mean of 10 replicates; bars represent the range of measurements. 

FIGURE 40. Numbers of bacteria or diatoms taken (organisms/mg h) by C. refulgens through grazing 
(first bar) or suspension feeding. All agglutinated material was removed from half of the attached foramini- 
fers (diagonal lines) while the remaining subpopulation was undisturbed (stippling) in order to determine 
if suspension feeding was facilitated by the agglutinated test extensions. Bar heights represent mean values 
(15 replicates for each suspension experiment and 60 replicates for the grazing experiment); error bars 
depict the range of values for each experiment. 



cytoplasm. Agglutinated tubes radiate away from the substrate and have been mea- 
sured to heights of five millimeters, and pseudopodia have been measured to extend 
an additional three millimeters from the tips of these tubes. The resulting canopy of 
branching tubes and pseudopodia potentially increases the volume of water available 
to suspension feeding by a factor of 10 to 20. Suspension feeding efficiency is further 
enhanced by the near proximity of other foraminifers and their tubes. 

Our experiments using radio-labeled prey demonstrated that C. refulgens cap- 



156 S. P. ALEXANDER AND T. E. DELACA 

TABLE I 
Free amino acu!> />.' />'< extrapallial space o/Adamussium colbecki 



Concentration of A A 

in extrapallial cavity Percent AA in 

nM 14 C protein hydrolysate* 





87.1 


9.3 


! ne 


165.5 


6.3 


Asparagine 


11.0 


0.0 


Glycine 


2066.3 


4.6 


Histidine 


0.0 


4.0 


Phenylalanine 


26.5 


6.7 


Proline 





5.6 


Serine 


76.8 


4.8 


Tyrosine 


11.8 


3.6 


Glutamic acid 


24.1 


11.8 


Valine 


13.1 


6.8 


Isoleucine 


8.9 


4.8 


Leucine 


12.7 


11.8 


Lysine 


17.3 


5.1 



* An analysis given by Amersham Corporation for its product CFB.25 ([U-' 4 C]algal protein hydroly- 
sate). 



tured both suspended diatoms and bacteria, and that capture efficiency was enhanced 
by agglutinated tubes by factors of 2.5 and 6. 1 for diatoms and bacteria, respectively. 
It is relevant to note here that to facilitate the experiment, concentrations of both 
suspended food sources were higher than the foraminifers would experience 
naturally. 

Several basic mechanisms of filter feeding are involved in systems such as that 
used by C. refulgens. According to Rubenstein and Koehl (1977), true sieving would 
not be an important contribution to particle capture in the nets of C. refulgens be- 
cause the average mesh size far exceeds the diameter of particles most likely to be 
encountered in the fine detritus of New Harbor. 'Direct interception' of particles by 
the sticky pseudopodia would form a large part of the filtering process, as would 
'motile particle deposition' of, for example, motile algae and bacteria. 'Gravitational 
deposition' of particles resuspended by the non-locomotory flapping motion of the 
scallops, is likely to provide an important source of captured material. The require- 
ment for water movement to allow filter feeding through the pseudopodial sieve must 
be satisfied almost entirely by movements of scallops, either locomotory (which oc- 
curs infrequently; see Mullineaux and DeLaca, 1984), non-locomotory, or by low 
velocity localized turbidity currents observed by DeLaca et al. (1980). 

How material is entrapped by pseudopodia of C. refulgens and transported to the 
agglutinated tubes or the cell body, has not been previously investigated. However, 
detailed information is available on the ultrastructural aspects of particle/prey entrap- 
ment and transport by pseudopodia of the Antarctic foraminifer Astrammina rara 
(see Bowser and DeLaca 1985a, b; Bowser et al., 1986), and Allogromia sp. (Bowser 
and McGee-Russell, 1982). In Allogromia sp. attachment is mediated by 'ultrami- 
crospikes' and 'ultramicrowebs' which possess special adhesive properties (Fig. 3, in 
Bowser and McGee-Russell, 1982; McGee-Russell et al., 1982). Structures almost 
identical to the ultramicrowebs of Allogromia sp. were observed in C. refulgens. These 
structures were found at the points of pseudopodial bifurcation, and in areas where 
particles were suspended (see Fig. 37) by the pseudopodia. That these structures sur- 



C. REFULGENS, MORPHOLOGY AND ECOLOGY 157 

vived the crude freeze-drying techniques available to us in Antarctica, without low 
calcium treatment and critical point drying, suggests that they may be even more 
extensive than our results indicate. These two taxonomically distant species appear 
to employ a similar mechanism for particle entrapment. 

Suspension feeding has been reported for a number of benthic foraminiferal gen- 
era, most of which possess elevated stalk-like tests anchored at one end to the sub- 
strate (See Christiansen, 1971; and Lipps, 1982; 1983 for reviews); in addition, two 
species of the benthic rotaliid Elphidium have been observed with three-dimensional 
pseudopodial networks extending into the seawater medium, which would be effi- 
cient collectors of free-floating food particles (Jepps, 1942; Sheehan and Banner, 
1972). In addition, the arborescent foraminifer Notodendrodes antarctikos DeLaca, 
living in New Harbor, captures bottom sediments which are brought into suspension 
by activities of larger benthic invertebrates (DeLaca el ai, 1980) and, as with C. reful- 
gens, this specialized mode of feeding is regarded as being an adaptation to an unusual 
oligotrophic environment. The availability of resuspended organic material could 
provide a more consistent source of food during the dark austral winter when benthic 
diatom productivity is low, and as such would compliment nutrients obtained by 
other mechanisms such as the uptake of free amino acids from the sediment and 
surrounding seawater in the case of N. antarctikos, and parasitism in the case of C. 
refulgens. 

The incorporation of agglutinated material into the test of calcareous foraminifera 
is an uncommon phenomenon which appears to be restricted to the suborder Milio- 
lina. Within the family Miliolidae, three genera (Sigmoilopsis, Ammomasilina, and 
Schlumbergerind) reportedly have test walls composed of agglutinated material 
bound by a calcareous cement; the genus Denstostomina has an external agglutinated 
layer of grains (Loeblich and Tappan, 1964). Within the Nubecularidae, Nubeculina 
has much coarse agglutinated material on the exterior of its chambers, and Nodobacu- 
laria incorporates occasional sand grains into the test (opera, cita.). However, among 
the suborder Rotaliina, test construction involving both agglutinated and calcareous 
material is rare. Nyholm ( 1 96 1 ) described a coniform stage of Cibicides lobatulus 
(which he regards as having developed from a zygote) with associated "tube-shaped 
structures composed of agglutinated material" extending vertically from the apex of 
the test, or occasionally, basally branching and leading from the aperture. Nyholm 
( 1 96 1 ) noted that an interspace of a few microns exists between the cytoplasm and 
the agglutinated wall of the coniform test; this may be morphologically analogous to 
the lumen of the circular agglutinated tube at the periphery of the test in C. refulgens. 
Indeed, regardless of which part of C. lobatulus ' life cycle these agglutinated tubes are 
associated, it is apparent that they are, in some respects, structurally similar. Nyholm 
(1961) did not suggest a function for the agglutinated tubes which project into the 
water above the attached sarcode (although he states that the agglutinated coniform 
test determines the form of the outer calcareous chambers), but in light of our findings 
with C. refulgens, it is tempting to speculate that they are also concerned with the 
deployment of pseudopodia into the surrounding water as a mechanism for suspen- 
sion feeding. 

The taxonomic and phylogenetic significance of extensive agglutinated additions 
to the tests of members of the genus Cibicides is not understood and remains an 
interesting question for taxonomists. 

ACKNOWLEDGMENTS 

This paper was made possible by the opportunity to use the SEM facility at the 
Department of Botany and Microbiology, University of Canterbury, Christchurch, 



158 S. P. ALEXANDER AND T. E. DELACA 

New Zealand, and e thank Mrs. K. Card for assistance. In addition, SEM work was 

performed .? /adsworth Center for Laboratories and Research, New York State 

Dept. of ) '.'>', and preparations were made in the laboratories of Dr. C. 

Rieder ' RR02157) with assistance from Dr. S. Bowser; the final manu- 

scrir neftted considerably from the comments of Dr. Bowser. Dr. S. M. Mc- 

:ridly provided resin-grinding facilities at the State University of New 

ny, for which we are grateful. Field and logistical support was provided 

National Science Foundation Division of Polar Programs, and research was 

upported by NSF grant DPP 83-05475. 

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Reference: Biol. Bull. 173: 160-168. (August, 1987) 



TIDAL H > GAMETOGENESIS: REPRODUCTIVE VARIATION 

G POPULATIONS OF GEUKENSIA DEMISSA* 

FRANCISCO J. BORRERO 

/ /artment of Biology and Belle W. Baruch Institute. University of South Carolina, 
Columbia, South Carolina 29208 

ABSTRACT 

High tidal populations of the mussel Geukensia demissa experience reduced filter 
feeding time as a result of aerial exposure. This study tested the hypothesis that such 
populations exhibit a temporal delay in their gametogenic cycle compared to popula- 
tions from the low intertidal. As predicted, quantitative estimations of gametogenic 
condition of mussels from 10 high tidal populations were lower than those of mussels 
from 1 1 low intertidal populations in May 1986. A two-fold difference in potential 
feeding time was accompanied by a delay of about two months in the reproductive 
activity of high tidal mussels. This study demonstrates that temporal reproductive 
variation among populations of (7. demissa across the intertidal zone may be as large, 
or larger than variation among latitudinally separated populations of this species. 
Site-to-site variation in timing of reproduction within the North Inlet Estuary may 
also be as large as temporal latitudinal variation. Level of occurrence in the intertidal 
zone and hence length of submersion and potential feeding time exert profound 
influence in the timing of the reproductive cycle of the ribbed mussel, Geukensia 
demissa. 

INTRODUCTION 

Variation in the timing of gametogenesis and spawning has been documented 
among populations of a number of marine invertebrates (Giese and Pearse, 1974, for 
a review). Such variation is common among populations of bivalve molluscs (Lubet 
et ai, 1981; Newell et al., 1982; Bayne and Newell, 1983), but the causes of this 
variability are not understood. Differences of environmental temperature (Orton, 
1920; Sastry, 1970) and seasonality of abundance and composition of food (Newell 
et al., 1982; Rodhouse et al., 1984) are thought to affect the reproductive cycles of 
marine bivalves. Latitudinally separated populations of the same species may exhibit 
large differences in the timing of reproductive activities (Sastry, 1970; Lubet et al., 
1981; Barber and Blake, 1983; Brown, 1984), but comparable variation also may be 
observed among populations separated by much smaller distances (Seed and Brown, 
1977; Newell et al., 1982). Microgeographic variation in the timing of reproductive 
activities may be due to site-specific differences in food supply (Bayne and Worrall, 
1980; Newell et al., 1982; Worrall et ai, 1983; Rodhouse et al., 1984). However, the 
causes of variation in timing and/or intensity of reproduction have been established 

Received 6 March 1987; accepted 1 April 1987. 
Abbreviation: GVF, Gamete Volume Fraction. 

* Contribution 679 from the Belle W. Baruch Institute for Marine Biology and Coastal Research, 
University of South Carolina. 

160 



TIDAL HEIGHT AND GAMETOGENESIS 161 

in only a few cases (Mann, 1 979; Velez and Epifanio, 1981; MacDonald and Thomp- 
son, 1986). 

Variation in height along the intertidal zone poses a dramatic gradient of food 
availability for filter feeding organisms. Since feeding can only occur during submer- 
sion, the potential feeding time is limited by the length of submersion. The effect of 
differences in potential feeding time on the energy balance of filter feeding animals is 
not clearly understood, but food availability and nutritional condition may strongly 
influence reproductive activity (Barber and Blake, 1983; Bayne and Newell, 1983). 
Therefore, differences in feeding time may have important consequences on the ga- 
metogenic cycles. 

This study examines the reproductive cycles of populations of the ribbed mussel 
Geukensia demissa (Dillwyn) (Bivalvia:Mytilidae) that occupy different tidal levels 
in the same salt marsh habitat. Previous studies suggested that diminishing the nutri- 
tional status of bivalve molluscs may delay their gametogenic cycle (Mann, 1979; 
Velez and Epifanio, 1981). Therefore, the specific hypothesis that the reproductive 
cycle of G. demissa will be delayed in high intertidal mussels which experience de- 
creased potential feeding time relative to low tidal mussels, was tested. 

Although G. demissa is a dominant secondary producer in salt marsh ecosystems 
of the eastern United States (Kuenzler, 1961; Fell et al., 1982; Jordan and Valiela, 
1982; Bertness, 1984), information on its reproductive biology is scarce. Ribbed mus- 
sel populations from New England and Connecticut exhibit a single spawning period 
from June through September (Brousseau, 1982; Jordan and Valiela, 1982), whereas 
in North Carolina and Georgia, peak reproductive activity might occur later, and/or 
gametogenic activity extend longer, in the year (McDougall, 1943; Kuenzler, 1961). 
This study provides a quantitative comparison of the reproductive cycles of popula- 
tions of G. demissa from South Carolina and describes temporal reproductive varia- 
tion associated with level of occurrence in the intertidal zone. 

MATERIALS AND METHODS 

During 1983-84, monthly samples of mussels were collected from a low tidal site 
(Site 1, Up Clambank) and a high tidal site (Site 2, Ely Creek), less than 1 km apart, 
within the North Inlet Estuary, South Carolina (Fig. 1 ). The high-tidal site is a short- 
form Spartina alt ern (flora marsh habitat, which is covered by water approximately 4 
hours per day. The low tidai site is an intertidal oyster (Crassostrea virginica) bed, 
where mussels occur among the oysters. This site is covered by water for about 8 
hours per day. Each sample consisted of approximately 25 mussels ranging from 25 
to 100 mm in shell length. 

Mussels were brought to the laboratory, scrubbed clean, and their shell lengths 
measured. A 1-cm 2 section was cut from the same area of the right mantle lobe of 
each animal and processed as follows. Serial sections 5 microns thick were prepared 
from paraffin embedded tissues and stained with hematoxylin and eosin. The frac- 
tional area of the mantle section that is composed of gametes (Gamete Volume Frac- 
tion, GVF ) was determined using stereology (Lowe et al, 1 982), and expressed as the 
mean GVF for each sample. GVF values were arcsine transformed to assure normal 
distribution. Two-way ANOVA (Sokal and Rohlf, 1981) was used to examine the 
effect of date and site on reproductive condition of mussels. The effects of date and 
sex were determined separately for each site, using two-way ANOVA. Mussels were 
separated into six shell length classes, and the effect of shell length on reproductive 
condition was determined using ANCOVA (Sokal and Rohlf, 1981). Mussel shell 
length was used as the covariate. Mean GVF values were detransformed for illustra- 



162 



F. J. BORRERO 




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FIGURE 1. Map of the North Inlet Estuary, South Carolina, indicating the ten sites studied. 1 = Up- 
Clambank, 2 = Bly Creek, 3 = Debidue Creek, 4 = Old Man Creek, 5 = Town Creek, 6 = Oyster Landing, 
7 = Oyster Island, 8 = Goat Island, 9 = Debidue Island, 10 = Jones Creek. 



tion in figures. Data on water temperatures at sites within the North Inlet were avail- 
able from the Long Term Ecological Research (LTER) program of the Baruch Insti- 
tute, University of South Carolina. 

RESULTS 

Temporal differences in the reproductive cycles of the high and low tidal popula- 
tions of G. demissa were observed. At the low tidal site, mussels had relatively con- 



TIDAL HEIGHT AND GAMETOGENESIS 



163 



QC 
U. 



UJ 

h- 

UJ 



O 



1.0 



0.8 



0.6 



0.2 



0.0 





30 



m 



TJ 

20 m 

DO 



10 



DO 

m 



O 



NDJFMAMJJASO 

1983 1984 



FIGURE 2. Reproductive cycles of Geukensia demissa during 1 983-84, from a low tidal population 
(Site I. Up-Clambank: ), and a high tidal population (Site 2, Ely Creek: O). Mean values of total GVF 
for approximately 25 individuals are plotted each month. Vertical lines indicate the standard error of the 
mean. Asterisks indicate significant differences (P < 0.05) between the two sites. Seasonal variation in 
surface water temperature () at Bread and Butter Creek (see Fig. 1). Each point represents a single mea- 
surement of temperature. 



stant and low GVF values from November to March, after which a rapid increase of 
GVF occurred, reaching a peak of about 0.57 by June (Fig. 2). By late April, no 
animals with sexually undifferentiated gonads were found. The increase of mean 
GVF closely followed the pattern of increasing water temperature in North Inlet, 
showing a significant (P < 0.05) and positive correlation (r = 0.776). This population 
maintained relatively high values of GVF for about four months (May to early Au- 
gust). The highest GVF value was observed in August (0.65), and spawning probably 
took place in this month and proceeded through September. It is possible that another 
spawning event of lesser intensity occurred at this site in June, which is supported by 
the observation of a significant (P < 0.05, Tukey a-posteriori test) decline of about 
20% in the mean GVF values between June and July. 

At the high tidal site, mussels exhibited a different reproductive cycle (Fig. 2). 
Gametogenesis started later, possibly between May and June. A rapid increase of 
GVF occurred between June and September, reaching an apparent peak of about 
0.65 in September. Spawning occurred in September, shortly after gametogenic de- 
velopment (Fig. 2). Since information on the reproductive condition of this popula- 
tion was available only from June to October, a correlation analysis between GVF 
and temperature is not very meaningful. However, the reproductive cycle of mussels 
at this site was delayed 2-3 months with respect to the pattern of increase in water 
temperature (Fig. 2). 

The ANOVA procedure indicated temporal differences (P < 0.05) in the repro- 
ductive cycle of the high and low tidal populations of Geukensia. The pattern of 
gametogenic development of male and female mussels was similar in the magnitude 
of the GVF values attained. However, male mussels appeared to start maturing earlier 
than females in both populations, since the sex ratio was skewed toward males in 



164 F J BORRERO 

samples colV n November and February at the low tidal site, and in June 

at the high U iie gametogenic cycle proceeded, the sex ratio did not differ 

signiJicar- later samples from either site. No evidence of hermaphrodit- 

j sm -, -d from 429 mussels examined. Sexually undifferentiated animals 

W ei' ^ April at the low tidal site, and until as late as July at the high tidal 

urther evidence for temporal displacement of the onset of gameto- 

:en the two populations. ANCOVA did not indicate a significant effect 

i length on the GVF values observed on animals from either population 

0.05). 

In summary, the reproductive cycles of the two populations differed in three ma- 
jor aspects: timing of the onset of gametogenesis; time of occurrence of spawning; 
and length of time mussels remained in a mature reproductive condition prior to 
spawning. Interestingly, despite the above temporal differences in reproduction, the 
highest GVF values observed on mussels from the two sites were similar (Fig. 2). 

A sampling program involving mussels from high and low tidal levels at a number 
of sites including the two original populations was conducted to determine whether 
the observed temporal differences in the reproductive cycles were restricted to the 
two sites studied, or whether they represented a general phenomenon among mussel 
populations across the intertidal zone. Since the 1983-84 samples indicated that the 
difference in reproductive condition was greatest at the end of the spring (Fig. 2), 
sampling was conducted on 24 May 1986. Based upon the results of the earlier sam- 
pling, the prediction that in May high tidal mussels should exhibit lower GVF values 
than mussels from the low intertidal was made. To test this hypothesis, ten sites were 
chosen such that low and high tidal mussel populations could be found (Fig. 1 ). Ap- 
proximately 15 mussels, 40-90 mm long, were collected from low and high tidal 
levels at each site. The high tidal level was Spartina marsh, similar at all sites, but the 
substrate at low tidal levels was marsh at some sites, and intertidal oyster beds at 
others. Therefore, the consistent difference among sites was tidal height and not habi- 
tat type. The samples were treated as described earlier. The effects of site, tidal level, 
and sex on reproductive condition were ascertained by ANOVA. Potential differ- 
ences in GVF among mussels of different sizes were examined using ANCOVA. 

The gametogenic condition of mussels from 10 sites in May 1986 is presented in 
Figure 3. Significant site-to-site variation in reproductive condition was observed (P 
< 0.0001, ANOVA). Despite this variation among sites, high tidal populations had 
lower GVF values than low tidal populations at all sites. These tidal-related differ- 
ences in gametogenic condition were highly significant (P < 0.0001, ANOVA). A 
Bonferoni procedure was used to assure 95% confidence in all statements regarding 
simultaneous pair-wise comparisons of GVF at each tidal level within sites. These 
comparisons indicate that the differences in GVF of high and low tidal mussels were 
significant in all but sites 4 (Old Man Creek) and 5 (Town Creek) (Figs. 1, 3). This 
analysis also suggests that tidal level has a larger effect on reproductive condition than 
does habitat type. All three habitat types were available at only one site (site 9, De- 
bidue Island). At this site, GVF values of low tidal marsh and low tidal oyster bed 
mussels were similar, and significantly different (P < 0.05) from that of high intertidal 
mussels (Fig. 3). The evidence for a larger effect of tidal level is further supported by 
the fact that regardless of habitat type, high tidal mussels had lower GVF values than 
mussels from the low intertidal at all sites. The observed differences in reproductive 
activity among tidal levels cannot be explained solely by temperature. Observations 
at the two primary sites indicate that no major temperature difference occurs between 
the two tidal levels, and that this difference is not systematic through time (Borrero 
and Hilbish, unpub. obs.). The temperature variation among sites within the estuary 
is also very small (LTER Data-base). 



TIDAL HEIGHT AND GAMETOGENESIS 



165 



1-0 r 



O 





^ 





\ 


i * 


0.8 


i * 


DC 

LL 

UJ 6 


A + J * 4 * 

? 1 


D 


I Jk 


- 1 0.4 

O 


'll ' 1* 

t\ ^ > 


W 0.2 


I \ * 


LU 

<r n n 


<J <} <j> 

1 1 1 1 1 I ^^^^ !H^^ J 



O 



123456 789 10 
SITE 



FIGURE 3. Reproductive condition of mussels from high and low tidal populations at ten sites in the 
North Inlet Estuary, on 24 May 1986. Mean values of total GVF for approximately 1 5 individuals and the 
standard error of the mean are plotted for each site and tidal level. Symbols represent mussels from high 
tidal marsh (O), low tidal marsh (), and low tidal oyster bed (A) substrates. The order of sites on the x- 
axis was established at random. Asterisks indicate significant (P < 0.05) differences between tidal levels at 
each site. 



Within the range of sizes considered, ANCOVA did not indicate a significant 
effect of mussel shell length on reproductive condition, and no difference was found 
in the reproductive condition of male and female mussels (P > 0.05, ANOVA). Ex- 
amination of the frequency of sexually active (male, female) versus undifferentiated 
mussels indicates again that the onset of gametogenesis is delayed at high tidal loca- 
tions, compared to populations from the lower intertidal. While all mussels were 
sexually active at most low tidal populations in May 1 986, undifferentiated animals 
were found at seven of the high tidal populations. The reproductive condition of 
mussels from the two original populations in the 1983-84 samples was very similar 
to that obtained from the same populations in 1986. 



DISCUSSION 

Current understanding of the feeding physiology of suspension-feeding bivalves 
indicates that the reduction in feeding time experienced by intertidal populations is 
directly proportional to the duration of aerial exposure. Bivalve molluscs exhibit a 
limited capacity to compensate for this reduction in feeding time (Bayne et al, in 
press). The reduction in energy intake by individuals at different tidal levels should 
be reflected in their productivity patterns. Tidal-related differences in productivity 
may result in variability of energy allocation to reproduction and timing of reproduc- 
tive activities. 

The results of this study demonstrate that level in the intertidal zone and hence 
length of submersion and potential feeding time affect the timing of the reproductive 
cycle of mussel populations. High tidal mussels in the North Inlet Estuary exhibited 



166 F. J. BORRERO 

delayed gonadal ;sc \ elopment compared to mussels from the lower intertidal. Spawn- 
ing was d ted only for the two populations from sites 1 and 2, but temporal 
difference onset of gametogenesis and reproductive condition of mussels in 
May i 9 e additional evidence that the delay in reproductive activity of high 
; is a general phenomenon. Studies on the reproduction of littoral 
1 other bivalve species indicate this may be a common pattern. Differ- 
>ductive maturity ofCardium edule from low intertidal and high shore 
? interpreted as due to the difference in synchrony of spawning at varying 
e levels (Boyden, 197 1 ). Spawning by intertidal Modiolus modiolus occurred be- 
tween late autumn and winter while it extended through most of the year in a subtidal 
population from Ireland (Seed and Brown, 1977). Hackney (1983) observed varia- 
tions in the timing of gonadal activity and spawning between well-flooded and irregu- 
larly flooded intertidal populations of Polymesoda caroliniana from Mississippi and 
Florida. Not surprisingly, this pattern seems to apply to other invertebrate groups. 
Palmer (1980) reported a temporal delay in the reproductive maxima of intertidal 
Microarthridion littorale (Copepoda), compared to a subtidal population at the same 
site. Similarly, the highest percentage of individuals with egg masses was associated 
with longer submergence time among two species of barnacles, interpreted as an effect 
of time available for feeding on reproductive activity (Page, 1984). 

A complete comparison of the effects of substrate type upon reproductive condi- 
tion cannot be achieved with the data from the present study. However, the effect of 
tidal height was apparent at all sites despite the heterogeneous nature of the substrates. 
This suggests that tidal height explains a major portion of the overall variance in 
reproductive condition. 

Microgeographic variation in the timing of reproductive activity documented in 
this study may be as great as that observed among latitudinally separated populations. 
This is evident from a comparison of the reproductive cycle of populations of 
G. demissa from the east coast of the United States. In Massachusetts, ripe mussels 
were observed in June-July, and spawning may occur in August-September (Jordan 
and Valiela, 1982). In Connecticut, gonadal development began in March, fully ripe 
individuals were observed June through September, and spawning occurred during 
the summer months (Brousseau, 1982). Spawning took place between August and 
September in North Carolina ( McDougall, 1 943 ), and Kuenzler (1961) indicated that 
spawning proceeded during July-August and into September in Georgia. No latitudi- 
nal pattern of variation appears to exist in the reproductive cycle of G. demissa. The 
results of the present study demonstrate temporal variation in reproductive activity 
within the North Inlet Estuary, as large as that reported from latitudinally separated 
localities. Furthermore, temporal variation in reproductive condition among tidal 
levels at a single site may be larger than variation between localities at greatly different 
latitudes. 

Similar results were obtained by Newell et al. (1982) in a study on reproductive 
variation ofMytilus edulis. A latitudinal pattern in the timing of reproductive activity 
of this species could not be established along the eastern coast of the United States. 
Site-to-site differences in food quantity and/or quality, and not temperature were 
identified as the major determinants of the timing of gametogenesis and spawning of 
Mytilus populations (Newell et al., 1982). The present study supports these conclu- 
sions. The two mussel populations described here differed two-fold in the time avail- 
able for feeding, and a similar difference in length of submersion applied for the high 
and low tidal populations at all sites sampled. 



TIDAL HEIGHT AND GAMETOGENESIS 167 

ACKNOWLEDGMENTS 

I thank Drs. T. J. Hilbish, F. J. Vernberg, D. Lincoln, S. A. Woodin, R. J. Feller, 
D. Edwards, D. S. Wethey, S. E. Stancyk, W. K. Michener, and three anonymous 
reviewers for discussion and criticism and for substantially improving the original 
manuscript with their critical reviews and comments. L. Barker provided water tem- 
perature information. Drs. R. I. E. Newell and V. S. Kennedy instructed on the use 
of stereology. Technical assistance by M. Walker and C. Cook is appreciated. Funds 
were provided by the Department of Biology and the Baruch Institute of the Univer- 
sity of South Carolina, a summer scholarship from the Southeast Chapter of the Ex- 
plorers Club, and the Aquaculture Fellowship from the South Carolina Wildlife and 
Marine Resources Division. 

LITERATURE CITED 

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(Lamarck) at its southern distributional limit. J. Exp. Mar. Biol. Ecol. 66: 247-256. 
BAYNE, B. L., ANDC. M. WORRALL. 1980. Growth and production of mussels Mytilus edulis from two 

populations. Mar. Ecol. Prog. Ser. 3: 317-328. 
BAYNE, B. L., AND R. C. NEWELL. 1983. Physiological energetics of marine molluscs. Pp. 407-5 15 in The 

Mollusca. Vol. 4, A. S. M. Saleuddin and K. M. Wilbur, eds. Academic Press, New York. 
BAYNE, B. L., A. J. S. HAWKINS, AND E. NAVARRO. Feeding and digestion in suspension feeding bivalve 

molluscs: the relevance of physiological compensations. Am. Zool. (in press). 
BERTNESS, M. D. 1 984. Ribbed mussels and Spartina alterniflora production in a New England salt marsh. 

Ecology 65(6): 1794-1807. 
BOYDEN, C. R. 1971. A comparative study of the reproductive cycles of the cockles Cerastoderma edule 

and C. glaucum. J. Mar. Biol. Assoc. U.K. 51: 605-622. 
BROUSSEAU, D. J. 1 982. Gametogenesis and spawning in a population ofGeukensia demissa (Pelecypoda: 

Mytilidae) from Westport, Connecticut. The Veliger 24(3): 247-25 1 . 
BROWN, R. A. 1984. Geographical variation in the reproduction of the horse mussel, Modiolus modiolus 

(Mollusca:Bivalvia). / Mar. Biol. Assoc. U.K. 64: 75 1-770. 
FELL, P. E., N. C. OLMSTEAD, E. CARLSON, W. JACOB, D. HITCHCOCK, ANDG. SILBER. 1982. Distribution 

and abundance of macroinvertebrates on certain Connecticut tidal marshes, with emphasis on 

dominant molluscs. Estuaries 99( 1 ): 2 1-28. 
GIESE, A. C., ANDJ. S. PEARSE. 1974. Introduction: general principles. Pp. 1-49 in Reproduction of Marine 

Invertebrates, Vol. 1, A. C. Giese and J. S. Pearse, eds. Academic Press, New York. 
HACKNEY, C. T. 1 983. A note on the reproductive season of the Carolina marsh clam Polymesoda carolini- 

ana (Bosc) in an irregularly flooded Mississippi marsh. Gulf Res. Rep. 7(3): 281-284. 
JORDAN, T. E., AND I. VALIELA. 1982. A nitrogen budget of the ribbed mussel, Geukensia demissa, and 

its significance in nitrogen flow in a New England salt marsh. Limnol. Oceanogr. 27( 1 ): 75-90. 
KUENZLER, E. J. 1961. Structure and energy flow of a mussel population in a Georgia salt marsh. Limnol. 

Oceanogr. 6: 191-204. 
LOWE, D. M., M. N. MOORE, AND B. L. BAYNE. 1982. Aspects of gametogenesis in the marine mussel 

Mytilus edulis L. J. Mar. Biol. Assoc. U.K. 62: 1 33-145. 
LTER Data-base. Long Term Ecological Research Program, Belle W. Baruch Institute for Marine Biology 

and Coastal Research, University of South Carolina, Columbia. F. J. Vernberg, Director. 
LUBET, P., J.-P. GIMAZANE, ANDG. PRUNUS. 1981. Etude du cycle de reproduction de Mytilus gallopro- 

vincialis (Lmk) (Moll. Lamellibranche) a la limite meridionale de son aire de repartition. Com- 

paraison avec les autres secteurs de cette aire. Haliotis 11:1 57- 1 70. 
MACDONALD, B. A., AND R. J. THOMPSON. 1986. Influence of temperature and food availability on the 

ecological energetics of the giant scallop Placopecten magellanicus. III. Physiological ecology, the 

gametogenic cycle and scope for growth. Mar. Biol. 93: 37-48. 
MANN, R. 1979. The effect of temperature on growth, physiology, and gametogenesis in the Manila clam, 

Tapes philippinarum (Adams & Reeve, 1 850). /. Exp. Mar. Biol. Ecol. 38: 1 2 1 - 1 33. 
McDouGALL, K. D. 1943. Sessile marine invertebrates of Beaufort, North Carolina. Ecol. Monogr. 13: 

321-374. 
NEWELL, R. I. E., T. J. HILBISH, R. K. KOEHN, AND C. J. NEWELL. 1982. Temporal variation in the 

reproductive cycle of Mvtilus edulis L. (Bivalvia, Mytilidae) from localities on the east coast of 

the United States. Biol. Bull. 162: 299-310. 



168 F. J- BORRERO 

ORTON, J. H. 1920. oerature, breeding and distribution in marine animals. J. Mar. Biol. Assoc. 

U.K. 1' K56. 

PAGE, H. variation in reproductive patterns of two species of intertidal barnacles, Pollicipes 

; ->y and Chthamalus fissus Darwin. /. Exp. Mar. Biol. Ecol. 74: 259-272. 
PA; anation in life-history patterns between intertidal and subtidal populations of the 

ae copepodMicroarthridion littorale. Mar. Biol. 60: 159-165. 

. C. M. RODEN, G. M. BRUNNELL, M. P. HENSEY, T. MCMAHON, B. OTTWAY, AND 
i. RYAN. 1984. Food resource, gametogenesis and growth ofMytilus edulis on the shore and 
-:pended culture: Killary Harbour, Ireland. J. Mar. Biol. Assoc. U.K. 64: 5 1 3-529. 
, A. N. 1970. Reproductive physiological variation in latitudinally separated populations of the 

bay scallop, Aequipecten irradians Lamarck. Bio. Bull. 138: 56-65. 

SEED, R., AND R. A. BROWN. 1977. A comparison of the reproductive cycles ofModiolus modiolus (L.), 
Ceraslodema (= Cardium) edule (L.), and Mytilus edulis L. in Strangford Lough, Northern Ire- 
land. OecologiaM: 173-188. 

SOKAL, R. R., AND F. J. ROHLF. 1 98 1 . Biometry, 2nd ed. W. H. Freeman and Co., New York. 859 pp. 
VELEZ, A., AND C. E. EPIFANIO. 198 1 . Effects of temperature and ration on gametogenesis and growth in 

the tropical mussel Perna perna (L.). Aquaculture 22: 2 1 -26. 

WORRALL, C. M., J. WIDDOWS, AND D. M. LOWE. 1 983. Physiological ecology of three populations of the 
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Reference: Biol. Bull. 173: 169-177. (August, 1987) 



DIFFERENCES IN THE DURATION OF EGG DIAPAUSE OF 

LABIDOCERA AESTIVA (COPEPODA: CALANOIDA) FROM THE 

WOODS HOLE, MASSACHUSETTS, REGION 

NANCY H. MARCUS* 
Woods Hole Oceanographic Institution, Woods Hole, Massachusetts 02543 

ABSTRACT 

The duration of diapause ofLabidocera aestiva eggs collected from the field and 
reared in the laboratory was determined at 5C. A clear seasonal trend was observed. 
Diapause eggs produced in the early fall required a much longer exposure to cold to 
yield a 50% hatch (CT 50 ) (i.e., the duration of diapause was longer) than eggs pro- 
duced later in the fall. Eggs produced by laboratory animals that were reared at 14C, 
8L-16D, required a shorter period of chilling to terminate diapause than the eggs of 
animals reared at 19C, 12L-12D. Considerable variation in the CT 50 value was also 
observed among laboratory cultures that were all reared under identical conditions, 
but which differed in terms of selection history. The results indicate that both the 
genotype of the egg and the conditions prevailing during oocyte formation influence 
the duration of diapause. Eggs that were stored at 5C for periods longer than 300 days 
no longer hatched upon warming. It is suggested that the variation in the duration of 
diapause is an adaptation that promotes synchronization of hatching by ensuring 
that all individuals terminate diapause at approximately the same time, and survival 
during the winter by conferring cold-hardiness. Synchronizing the onset of post-dia- 
pause development is also discussed as an alternative mechanism for achieving syn- 
chronous hatching. 

INTRODUCTION 

The calanoid copepod, Labidocera aestiva, is a seasonal member (summer and 
fall) of the planktonic community in the Woods Hole region. In this area most 
L. aestiva females have the genetic potential to produce two types of eggs: subitaneous 
and diapause (Marcus, 1982). Subitaneous eggs are produced during the summer 
and fall; diapause eggs are produced during the fall. Both egg types begin to develop 
following their release by females. Subitaneous eggs typically hatch within 1 to 4 days 
at 2 1 to 23C (Marcus, 1 979). Diapause eggs enter a refractory phase after 24 to 48 h 
of development. During the refractory phase, further embryogenesis is not apparent 
(Marcus, pers. obs.) and diapause eggs cannot be induced to hatch even if conditions 
are favorable. The duration of diapause (i.e., the length of the refractory period) is 
positively related to the temperature at which eggs are held (Grice and Gibson, 1975; 
Marcus, 1979). Once the refractory phase is completed, post-diapause development 
and hatching occurs if conditions (e.g., temperature) are favorable. For instance, at 
21 to 23C hatching typically occurs within 1 to 2 days (Marcus, 1979). Several field 
and laboratory studies (Marcus, 1979, 1980, 1 984) support the claim that the perpet- 



Received 2 March 1987; accepted 14 May 1987. 

* Present address: Department of Oceanography, Florida State University, Tallahassee, FL 32306. 

169 



170 N. H. MARCUS 

uation of L. year after year in the Woods Hole region is due to the diapause 

eggs whid rater on the sea-bottom and hatch in the spring. 

:-A~td laboratory studies on L. aestiva I observed that many diapause 
egj-, <_d to hatch at 19C following a chilling period of 4 weeks at 5C. 

eggs would hatch with a shorter period of chilling while others re- 
n longer exposure to cold. A comparison of diapause eggs obtained from 
collected in the field showed that the period of chilling that would result in a 
hatch at 19C was longer for the diapause eggs of females collected early in the 
fail (Marcus, 1986). This study examines in more detail the seasonal variation in the 
duration of diapause of eggs of freshly caught animals from the field and compares 
the results to values obtained for diapause eggs of females reared in the laboratory. 
The results indicate that the genotype of an egg and the environmental factors acting 
during oocyte formation influence the duration of diapause. Based on the results, I 
suggest that the variation in the duration of diapause is an adaptation that promotes 
synchronization of hatching in the field by ensuring that all individuals terminate 
diapause at approximately the same time, and survival during the winter by confer- 
ring cold-hardiness. 

MATERIALS AND METHODS 

Diapause eggs were obtained from animals collected at one to two week intervals 
over a period of 2 years from October 1 98 1 to October 1 983. For each sampling date, 
adult females were collected from Vineyard Sound by towing a 3 / m diameter, 243 ^ 
mesh plankton net for 10 min. Water temperature was determined on a surficial 
bucket sample for all but two collection dates. For these dates, water temperature 
was estimated based on the daily temperature record for water off the Woods Hole 
Oceanographic Institution dock. A comparison of several dates showed that the 
WHOI values were typically about 1 C less than Vineyard Sound values. Field sam- 
pling dates and surface water temperature at the time of collection are shown in Table 
I. In the laboratory females were transferred to 100 ml dishes containing 5 /im-filtered 
seawater and the dinoflagellate Gymnodinium nelsoni (500 cells/ml). The dishes were 
incubated overnight at 19C. The next day eggs were collected by pipette, pooled in 
a separate dish of filtered seawater, and returned to the incubator for 2-4 days to 
allow the subitaneous eggs to hatch. Unhatched eggs that appeared to be diapause 
eggs (i.e., the interiors were green, with a clear perimeter) were distributed into 75 ml 
glass screw capped jars (20-25 eggs/jar) containing filtered seawater, and refrigerated 
at 5C. Eggs that were obviously non-viable (i.e., the interiors were brownish, granu- 
lar, and disintegrating) were discarded. Every 2 to 4 days, a jar was removed (except 
for the 2 collections in 1981 for which duplicate jars were removed), warmed to 1 9C, 
and held at that temperature. After 4 to 5 days the proportion of hatched eggs was 
ascertained. The average hatch of the duplicates was recorded for the two 1 98 1 collec- 
tion dates. 

Eggs of laboratory-reared animals were from 1 2 different cultures that were reared 
either at a temperature of 14 or 19C (1C), and a photoperiodic regimen of 8L- 
1 6D or 1 2L- 1 2D. The eggs collected from each culture were 1 -2 days old. The adults 
were approximately 2 weeks past reproductive maturity. The cultures represented 
specific generations of three inbred lines that were being perpetuated as part of a long- 
term selection experiment designed to assess the potential for evolutionary change in 
the diapause response threshold (i.e., the necessary conditions for the expression of 
diapause). Each line was initiated from 500-1000 nauplii that were derived from 
pooled batches of either subitaneous or diapause eggs produced by 60 females col- 



DURATION OF EGG DIAPAUSE 171 

TABLE I 

Collection dates, surface water temperature (C). chilling lime (days) required for initial hatch, CT SO values 
(days), and regression parameters pertaining to diapause eggs of field collected females 

Date C Initial CT 50 r 2 Slope 



9/20/82 


19 


14 


23.19 


.88 


7.31 


9/26/83 


21 


18 


28.82 


.80 


9.07 


9/27/82 


19 


12 


28.61 


.79 


4.14 


10/04/82 


18 


14 


27.40 


.88 


6.03 


10/11/83 


18 


23 


26.13 


.90 


13.69 


10/12/82 


17 


16 


23.02 


.78 


10.11 


10/19/81 


14 


4 


15.44 


.89 


3.66 


10/21/82 


14 


6 


24.14 


.74 


3.71 


10/24/83 


16 


20 


22.37 


.84 


8.61 


11/01/82 


14 


8 


17.30 


.91 


5.25 


11/10/82 


13 


6 


16.53 


.90 


4.81 


11/23/82 


11 


4 


15.36 


.80 


3.22 


11/29/82 


10 


2 


8.98 


.89 


3.44 


12/01/81 


7 


6 


10.99 


.99 


3.16 


12/08/82 


10 


2 


10.17 


.96 


2.71 



lected from the field. Two of the lines were termed subitaneous. Each generation was 
perpetuated from 500-1000 nauplii derived from just the subitaneous eggs that were 
produced by the preceding generation of animals. A third diapause line was perpetu- 
ated in a similar manner, but from just diapause eggs. The specific rearing conditions 
and selection histories are shown in Table II. The proportion of subitaneous and 
diapause eggs produced by each generation of animals varied within and between the 
lines. The diet for all cultures consisted of a standard mix of four dinoflagellates. 
General methods for rearing of L. aestiva have been described previously (Marcus, 
1980). Eggs from each culture were incubated at 19C for 4 to 5 days after which 
the diapause eggs were distributed (20 to 30 eggs/jar) into 75 ml jars. The jars were 
refrigerated at 5C. At 2 to 4 day intervals the jars were removed, warmed to 19C, 
and held at that temperture. The proportion of eggs that hatched after 4 to 5 days was 
determined. 

For each field sampling date and laboratory culture, values of percent hatch were 
transformed to probit values (Finney, 1952). A regression analysis was performed 
with these values versus the number of days chilled (Iog 10 ) to derive an estimate of 
the days of chilling required to promote a 50% hatch (CT 50 ). Calculations were done 
with an IBM PC and the statistical software package, STATPRO. 

The effect of long-term storage at 5C on egg viability and hatching was examined 
for the 2 sets of diapause eggs obtained from culture 339. After the initial analysis 
period, jars of eggs were removed at intervals of up to 4 weeks for more than a year. 
The hatch of these eggs after warming to 1 9C was ascertained as described above. 

RESULTS 

In general, a shorter period of chilling was necessary to promote initial and 50% 
hatching of eggs produced by females collected from the field later in the fall (Table 
I, Fig. 1 ). The results of the Probit transformation and regression analysis permit a 
quantified comparison of these differences and the derivation of the median effective 
chilling period (i.e., the number of days of chilling that promote a 50% hatch). The 



172 



N. H. MARCUS 



, -s. 

- 

E 
N 
T 

H 
A 

T 
C 
H 



lOOf 



75" 



50" 



25" 



o-Lt 




DAYS CHILLED 

FIGURE 1. Percent hatch of diapause eggs, from field-collected females, at 19C after chilling at 5C 
for the designated number of days. Each set of connected points represents a specific sampling date. Dates 
for each month are grouped by the indicated symbols. 



regression parameters (slope, r 2 ) and CT 50 values are shown in Table I. The coefficient 
of determination values (r 2 ) ranged from .74 to .99 indicating that the linear regres- 
sion relationship was a good one for estimating the CT 50 . For the diapause eggs of 
field-collected females, the CT 50 values ranged from 8.98 to 28.82 days. The slope 
values of the regression ranged from 2.71 to 13.69 probit value/days (log, ). This 
latter parameter provides an indication of the time spread of diapause duration 
around the median. A high value corresponds to a very short interval for the time 
from initial to maximal hatching. The highest values tended to occur during Septem- 
ber and October, and the lowest during November and December. This same pattern 
was found for the CT 50 values. Further analysis revealed that a very good positive 
correlation (r 2 = .84) existed between CT 50 values and surface water temperature at 
the time of sampling (Fig. 2). 

The median effective duration of chilling also differed among the laboratory 
reared groups although the range of values was not as great as observed for the field 
group. The regression parameters (slope, r 2 ) and CT 50 values are shown in Table II. 
As for the field group the r 2 values were high (.71 to .94). The CT 50 values ranged 
from 5.82 to 21.09 days. The slope values of the regression ranged from 1 .70 to 5.23 
probit value/days (log, ). The lowest CT 50 values were obtained for the 3 cultures 
(370, 371, 374) that were reared at 14C and 8L-16D. For 2 pairs of cultures, 370 
and 372, and 371 and 373, the within pair cultures were established from the same 
pool of eggs in the 23rd generation, but were reared at the two alternative sets of 
conditions. In both cases the CT 50 values were lower for the cultures reared at 14C, 
and 8L-16D. A third unpaired culture (374) was reared at 14C and 8L-16D and 
also yielded the third lowest CT 50 value. 

The two sets of eggs that were obtained from culture 339 were collected on differ- 
ent days and the CT 50 values differed by almost 5 days. The long term response to 
chilling was also different for the two sets (Fig. 3). The hatch after chilling increased 
more rapidly during the first 30 days for 339b, but a high hatch was maintained for 



DURATION OF EGG DIAPAUSE 



173 



CT50 VALUES VS FIELD TEMP. 



30" 



C 

T 

5 




20" 




10" 







5 10 15 20 

TEMPERATURE ( C) 



FIGURE 2. Linear regression analysis of CT 50 value of each field sample and the surface water temper- 
ature at the time of collection. 

only 150 to 200 days whereas a high hatch was maintained by the eggs of 339a for 
almost 300 days. Although the hatch of both sets dropped off to near 0% levels after 
300 days, many of the eggs in both sets still appeared viable. 

DISCUSSION 

This study shows that the median effective number of days of chilling at 5C de- 
creased as the fall season progressed for the diapause eggs of field-collected females. 



TABLE II 

Culture # (generation, subitaneous-s or diapause-d line), rearing conditions (photoperiod and 
temperature), CT 50 values (days), and regression parameters pertaining 
to diapause eggs of laboratory-reared females 



Culture # 



Conditions 



CT 



50 



* Eggs collected from same culture, but on two different days. 



Slope 



370 (23s) 


14C, 8L-16D 


10.43 


.85 


2.18 


371 (23s) 


14C, 8L-16D 


5.82 


.71 


1.70 


374 ( 6s) 


14C, 8L-16D 


10.74 


.90 


2.92 


375 ( 3s) 


19C, 12L-12D 


12.16 


.91 


2.59 


372 (23s) 


19C. 12L-12D 


11.77 


.78 


3.11 


373 (23s) 


19C, 12L-12D 


14.33 


.74 


2.37 


376 ( 7d) 


19C, 12L-12D 


19.33 


.78 


2.41 


323 ( 7s) 


19C 12L-12D 


17.92 


.83 


4.42 


329 (15s) 


19C, 12L-12D 


13.64 


.87 


3.04 


325 ( 4d) 


19C 12L-12D 


21.09 


.78 


3.27 


339a( 5d)* 


19C, 12L-12D 


16.59 


.93 


5.23 


339b( 5d)* 


19C, 12L-12D 


11.81 


.94 


3.73 


330 (15s) 


19C, 8L-16D 


15.66 


.84 


3.54 



174 



N. H. MARCUS 



E 
R 

C 

N 
T 

H 
A 
T 
C 
H 



100- 



X: 339a 
: 339b 




100 



200 



300 



400 



DAYS CHILLED 



FIGURE 3. Percent hatch of diapause eggs, produced by culture 339 on different days (a and b), at 
19C after chilling at 5C for the designated number of days. 



The range of values spanned 20 days. Similar seasonal trends have been reported for 
the diapause stages of insects (Burdick, 1937; Church and Salt, 1952). During the 
collecting period of L. aestiva, water temperatures ranged from 20.5 to 6.9C, a 
difference of approximately 14.0C. The two temperatures at which the laboratory 
animals were reared differed by 5C and the number of days of chilling required to 
achieve a 50% hatch differed by as much as 15.5 days. At a constant 19C, the range 
in CT 50 values was about 10 days for the eggs of laboratory-reared animals. Thus for 
the laboratory-reared animals considerable variation in the median effective days of 
chilling was obtained, despite the fact that the environmental conditions were the 
same. This variation must reflect genetic differences. Further evidence for genetic 
variation are the different responses observed for the eggs of culture 339 that were 
collected on different days. Since the eggs all came from the same culture, the only 
possible explanation is that the eggs collected on the different days were produced by 
different mixes of parents. Although the cultures of animals that were used for the 
analyses represented different generations of three genetically distinct lines, no obvi- 
ous association was observed between diapause duration and generation number or 
selection history. 

Although genetic differences appear to be important, environmental factors may 
also have an effect. The three shortest times to achieve a 50% hatch were obtained for 
the cultures that had been reared at 14C and 8L-16D (Table II). This same relation- 
ship with temperature and short-daylengths was evident for the eggs of the field col- 
lected animals. The work of Denlinger and Bradfield (1981) on the tobacco horn- 
worm provides a possible explanation for these trends. They showed that the duration 
of diapause was influenced by the number of short day cycles perceived by individu- 
als. As the number of short day cycles experienced by an individual increased, the 
duration of diapause decreased. They concluded that in the field the duration of dia- 
pause is shorter for individuals entering diapause late in the fall because declining 
temperatures lead to slower development and therefore a longer exposure to short 
daylengths. If this mechanism characterizes L. aestiva it is unlikely that the oocyte 



DURATION OF EGG DIAPAUSE 175 

itself could perceive the number of short day cycles. Hence, the effect would have to 
be mediated through the parent female as a "maternal effect." In the case of L. ae- 
stiva, it is also possible that declining temperatures directly influence the physiologi- 
cal state of the female and, in turn, oogenesis. The differences observed for the eggs 
of field-collected animals could also result from variation in maternal age. Animals 
collected late in the fall might be older than ones collected early in the fall. However, 
this would not explain the variation expressed by the eggs of laboratory-reared fe- 
males since all of the animals were similar in age. Krysan and Branson (1977) con- 
ducted specific crosses with the corn rootworm and showed that the duration of dia- 
pause was affected both by the genotype of the embryo and a maternal component. 
Further experiments are needed to assess the relative importance of these compo- 
nents in L. aestiva. 

The long term response to chilling observed for the eggs of culture 339 (Fig. 3) is 
very similar to patterns observed for several insect species (Hussey, 1955; Cranham, 
1972; Lees, 1955). For these species, the percent of individuals terminating diapause 
increased to a maximum with increasing length of exposure to cold, then remained 
high with longer exposure to cold, and finally declined with excessive time of expo- 
sure to cold. In each case the percent terminating diapause remained low after exces- 
sive exposure, but the interpretation of the results differed among authors. Hussey 
(1955) suggested that the decline was part of an annual cycle and that given enough 
time the percent terminating diapause would increase again. Hussey believed that 
there was an internal gating rhythm that controlled emergence from diapause. How- 
ever, since he did not carry the experiments through for another year it cannot be 
certain that death had not occurred. The rhythm concept was not discussed by the 
others despite similar results. They concluded that the eggs had lost viability and 
would never hatch. It would certainly be advantageous for an egg of L. aestiva to 
remain viable beyond one season. L. aestiva eggs that are buried do not hatch despite 
favorable temperatures (Marcus and Schmidt-Gengenbach, 1986). The probability 
of completing development should be higher for a diapause egg (from the previous 
fall) that is uncovered no later than the summer than for an egg which is not uncov- 
ered until October or November. If hatching occurred only in October and Novem- 
ber, the likelihood of completing development should be diminished due to declining 
temperatures. A gating rhythm that controlled the onset of post-diapause develop- 
ment would reduce the probability of eggs hatching at an inappropriate time. How- 
ever, this study does not support such an hypothesis. After 300 days of chilling, the 
percent hatch of L. aestiva eggs after warming declined. After more than 400 days of 
chilling, hatching has not increased again though many of the eggs look viable. Thus 
this study indicates that eggs cannot survive more than 300 days of constant exposure 
to 5C in the laboratory. However, this life span may be quite different in the field, 
where eggs probably experience long periods of anoxia and exposure to hydrogen 
sulfide. Although the effect of such parameters on the viability of L. aestiva eggs is 
not known, it has been reported (Uye et al, 1984) that exposure to organic pollution 
reduces the viability of resting eggs of neritic marine copepods. 

From studies of insects and freshwater copepods, I suggested (Marcus, 1979) that 
L. aestiva eggs terminate diapause at different times during the winter in the field, 
and are held at a stage of pre-hatch readiness because water temperatures are below 
the threshold for post-diapause development and hatching. This study supports that 
hypothesis, although the duration of diapause under field conditions appears to differ 
from that observed in the laboratory where temperature was held constant at 5C. 
The data (Marcus, 1984) for eggs collected from bottom sediments in Buzzards Bay, 
Massachusetts, indicated that the refractory phase was not completed by all eggs in 



176 N. H. MARCUS 

December. It was suggested that the eggs which failed to hatch at this time were pro- 
duced late 2 fail and had not completed the refractory period. By February all 
eggs apix ve completed the refractory phase as evidenced by the high hatch 
of eggs i red at 19C. The present study does not support that suggestion. Be- 
cau c' sonai variation in the duration of diapause it is possible that eggs 
cted from sediments in December and did not hatch in the labora- 
warrning, were produced in September as well as in December, 
i ature is not the only environmental parameter that affects the transition 
m diapause to development. Hatching is also affected by light and oxygen concen- 
tration (reviewed by Grice and Marcus, 198 1). Although the effect of these parame- 
ters on the termination of diapause and the onset of post-diapause development in 
marine copepods has not been clarified. Brewer (1964) reported that exposure to re- 
duced oxygen concentrations was necessary to terminate egg diapause in the freshwa- 
ter copepod, Diaptomus stagnalis, and Watson and Smallman ( 197 1) suggested that 
photoperiod was a necessary cue for the resumption of development in Diacyclops 
navus. The transition from dormancy to development is mediated by pH in brine 
shrimp (Busa and Crowe, 1983). 

Diapause is an important factor in the synchronization of life cycles (Tauber et 
al, 1986). Two possible ways in which synchronization can be achieved are synchro- 
nizing the termination of diapause and synchronizing the onset of post-diapause de- 
velopment and hatching. Both mechanisms characterize L. aestiva in Woods Hole 
waters. Since diapause eggs are produced over a span of several months, the longer 
diapause of eggs produced early in the fall ensures that they do not terminate diapause 
until winter temperatures have declined below the threshold for post-diapause devel- 
opment. Conversely, the shorter diapause of eggs produced late in the fall ensures 
that diapause will be completed prior to the time when water temperature exceeds 
the threshold in the spring. Since diapause in the field terminates by February or 
March in Woods Hole waters (Marcus, 1 984) and hatching does not occur until May 
(Grice and Gibson, 1975) the precise coincidence of diapause termination should be 
less important than the coincidence of the onset of post-diapause development in the 
promotion of synchronous hatching. Thus, as long as the refractory phase is com- 
pleted before the threshold for post-diapause development or hatching is exceeded, 
synchronization of hatching should still occur. The coincidence of diapause termina- 
tion among overwintering eggs may be more critical at more southern latitudes where 
water temperatures at the time of diapause termination are more likely to exceed the 
threshold and thus individuals would resume development as soon as the refractory 
period ended. The results also suggest that diapause is important because it promotes 
the survival of individuals by conferring cold-hardiness. Diapause eggs which com- 
plete their refractory period by January (Marcus, 1 984), can tolerate exposure to cold 
winter temperatures. However, subitaneous eggs do not survive extended exposure 
to such temperatures (Grice, unpub.). I suggest that the variation in diapause duration 
expressed by L. aestiva is an adaptation that promotes synchronization by ensuring 
that all individuals terminate diapause at approximately the same time, and survival 
by conferring cold-hardiness. 

ACKNOWLEDGMENTS 

I thank C. Fuller and P. Alatalo for their valuable assistance in the field and labora- 
tory. J. Schmidt-Gengenbach, S. Twombly, and two anonymous reviewers provided 
helpful criticism of the manuscript. Supported by NSF Grants OCE82- 14882 and 
OCE85-09863. 



DURATION OF EGG DIAPAUSE 177 

LITERATURE CITED 

BREWER, R. H. 1 964. The phenology ofDiaptomus stagnalis (Copepoda:Calanoida): the development and 

the hatching of the eggs stage. Physiol. Zoo/. 37: 1-20. 
BURDICK, H. 1937. The effects of exposure to low temperature on the developmental time of embryos of 

the grasshopper Melanoplus differentially (Orthoptera). Physiol. Zoo/. 10: 156-170. 
BUSA, W., AND J. CROWE. 1983. Intracellular pH regulates transitions between dormancy and develop- 
ment of brine shrimp (Anemia salina) embryos. Science 221: 366-368. 
CHURCH, N., AND R. SALT. 1952. Some effects of temperature on development and diapause in eggs of 

Melanoplus bivittatus (Say) (Orthoptera: Acrididae). Can. J. Zoo/. 30: 99-1 7 1 . 
CRANHAM, J. 1972. Influence of temperature on hatching of winter eggs of fruit-tree red spider mite, 

Panonychus ulmi (Koch). Ann. App. Biol. 70: 1 19-1 37. 

DENLINGER, D., AND J. BRADFIELD. 1981. Duration of pupal diapause in the tobacco hornworm is deter- 
mined by number of short days received by the larva. /. Exp. Biol. 91: 33 1-337. 
FINNEY, D. 1952. Probit Analysis: A Statistical Treatment of the Sigmoid Response Curve. Cambridge 

University Press, London. 
GRICE, G., AND V. GIBSON. 1975. Occurrence, viability, and significance of resting eggs of the calanoid 

copepod Labidocera aestiva. Mar. Biol. 31: 335-337. 
GRICE, G., AND N. MARCUS. 1981. Dormant eggs of marine copepods. Oceanogr. Mar. Biol. Ann. Rev. 

19: 125-140. 
HUSSEY, N. 1955. The life histories of M^ay//gw/s;?mHO/ro/7/n/.9 Wachtel(Hymenoptera:Chalcidoidea) 

and its principal parasite with descriptions of the developmental stages. Trans. R. Entomol. Soc. 

Lond. 106: 133-151. 
KRYSAN, J., ANDT. BRANSON. 1977. Inheritance of diapause intensity in Diabrotica virgifera. J. Hered. 

68:415-417. 
LEES, A. 1953. Environmental factors controlling the evocation and termination of diapause in the fruit 

tree red spider mite Metatetranychus ulmi Koch (Acarina:Tetranychidae). Ann. Appl. Biol. 40: 

449-486. 
MARCUS, N. 1979. On the population biology and nature of diapause of Labidocera aestiva (Copepoda: 

Calanoida). Biol. Bull. 157: 297-305. 
MARCUS, N. 1980. Photoperiodic control of diapause in the marine calanoid copepod Labidocera aestiva. 

Biol. Bull. 158:311-318. 
MARCUS, N. 1982. The reversibility of subiatneous and diapause egg production by individual females of 

Labidocera aestiva (Copepoda:Calanoida). Biol. Bull. 162: 39-44. 
MARCUS, N. 1984. Recruitment of copepod nauplii into the plankton: importance of diapause eggs and 

benthic processes. Mar. Ecol. Prog. Ser. 15: 47-54. 
MARCUS, N. 1986. Population dynamics of marine copepods: the importance of photoperiodism. Am. 

Zoo/. 26: 469-477. 
MARCUS, N., AND!. SCHMIDT-GENGENBACH. 1986. Recruitment of individuals into the plankton: the 

importance of bioturbation. Limnol. Oceanogr. 31: 206-210. 
TAUBER, M., C. TAUBER, AND S. MASAKI. 1 986. Seasonal Adaptations of Insects. Oxford University Press, 

New York. 
UYE, S., M. YOSHIYA, K. UEDA, ANDS. KASAHARA. 1984. The effect of organic sea-bottom pollution on 

survivability of resting eggs of neritic calanoids. Crustacean Suppl. 1: 390-403. 
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Reference: Biol. Bull. 173: 178-187. (August, 1987) 



'HEMICAL FEATURES OF SHRIMP HEMOCYTES 

JO ELLEN HOSE, GARY G. MARTIN, VAN ANH NGUYEN, 
JOHN LUCAS, AND TEDD ROSENSTEIN 

Department of Biology. Occidental College, Los Angeles, California 90041 

ABSTRACT 

Morphological studies suggest that there are several types of decapod hemocytes; 
however, distinguishing criteria based on conventional staining techniques are often 
subtle or ambiguous. Cytochemical features of ridgeback prawn (Penaeidae: Sicyonia 
ingentis) hemocytes were studied using specific stains for lysosomes, cytoplasmic con- 
tents, and granule enzymes. This approach facilitates the differentiation of cell types 
in the ridgeback prawn and provides information on the functions of and relation- 
ships among different cell types. 

Agranular hemocytes and a subgroup of small granule hemocytes contain exten- 
sive cytoplasmic glycoprotein deposits which display smudgy, intense staining with 
Sudan black B. As previously shown, coagulogen the clotting material in deca- 
pods stains with Sudan black B when extracted from lysed hemocytes. Other hemo- 
cyte types display light staining limited to granule membranes. 

Lysosomes are not observed in agranular cells and are rarely present in small 
granule hemocytes with glycoprotein deposits. Small granule hemocytes without de- 
posits and large granule hemocytes contain numerous lysosomes as shown by the 
presence of acid phosphatase, /3-glucuronidase, and nonspecific esterase. Acid phos- 
phatase is observed in the Golgi body of these cells, within small vesicles, and in 
small granules. The granules in large granule hemocytes rarely show acid phosphatase 
reaction, yet small acid phosphatase-positive vesicles fuse with the large granules. The 
acid phosphatase in the large granules may exist in an inactive form. Prophenoloxi- 
dase activity is localized only in large granules. The physiological significance of he- 
mocyte cytochemistry is also discussed. 

INTRODUCTION 

Crustacean hemocytes perform a variety of physiological and pathological func- 
tions including coagulation (Ravindranath, 1980), phagocytosis, recognition of for- 
eign material, carbohydrate transport, and encapsulation (Bauchau, 1981). Several 
hemocyte categories have been recognized in decapod crustaceans based on morpho- 
logical criteria. Morphological features are often subtle and ambiguous, and are not 
readily recognized by other investigators. In addition, morphological criteria are 
rarely based on properties that facilitate the differentiation between stages in hemo- 
cyte maturation or among cells with different physiological functions. Hence, previ- 
ous investigations have failed to define a clear correspondence between various cell 
types and their functions. 

In an attempt to develop a comprehensive description of crustacean hemocyte 
formation and function, cytochemical techniques were used to complement our pre- 
vious morphological description of shrimp hematopoietic tissue (Martin et al, 1987) 
and circulating hemocytes (Martin and Graves, 1985). Electron microscopic exami- 

Received 2 January 1987; accepted 19 May 1987. 

178 



CYTOCHEMISTRY OF SHRIMP HEMOCYTES 179 

nation of hemocytes from the ridgeback prawn (Penaeidae: Sicyonia ingentis) shows 
the existence of four cell types: agranular, small granule with cytoplasmic deposits, 
small granule without cytoplasmic deposits, and large granule hemocytes. Agranular 
hemocytes are small cells with a high nucleus:cytoplasm ratio (Martin et al, 1987). 
Their cytoplasm contains little other than aggregations of electron-dense deposits. A 
subset of small granule hemocytes contains similar electron-dense cytoplasmic de- 
posits, one to six round striated granules, and occasional electron-dense granules. In 
contrast, a distinct subset of small granule hemocytes and the large granule hemocytes 
lack cytoplasmic deposits and striated granules. These hemocytes have many (> 10) 
electron-dense, electron-lucent, or punctate granules which range in diameter from 
0.4 nm in small granule hemocytes to 0.8 ^m in large granule cells. Intermediate 
stages were observed between agranular hemocytes and small granule hemocytes with 
deposits and between small granule hemocytes without deposits and large granule 
hemocytes, suggesting the existence of two distinct hemocyte lines. 

In view of the difficulty in accurately identifying certain hemocyte categories at 
the light microscope level, various enzymatic and cytochemical methods were evalu- 
ated for use in hemocyte classification. The goals of this study are to (1) identify 
cytochemical stains which can be used to differentiate specific hemocyte types, and 
(2) provide useful information on the function of the various cell types. 

MATERIALS AND METHODS 

Animals 

Ridgeback prawns were collected and maintained as previously described (Martin 
et al., 1987). Shrimp averaged 14.5 g and were in molt stages C and D (Ander- 
son, 1985). 

Tissue collection and preparation 

Hemolymph (usually 0.2 cc) was withdrawn from the ventral sinus or heart into 
a 1 cc syringe containing anticoagulant (Martin and Graves, 1985). Hemocyte smears 
were then prepared on glass microscope slides, allowed to air dry, and used for light 
microscopy. 

Hemocytes and hematopoietic nodules to be examined at the electron micro- 
scopic (EM) level were fixed in 2.5% glutaraldehyde in 0. 1 M sodium cacodylate (pH 
7.8) containing 12% glucose for 1 h at room temperature. Following a 30 min wash 
in 0.1 M sodium cacodylate (pH 7.8) containing 24% sucrose, the tissues were post- 
fixed in 1% OsO 4 in 0.1 M sodium cacodylate for 1 h at room temperature, dehy- 
drated in a graded series of ethanol, and infiltrated and embedded in Spurrs' (1969) 
low viscosity plastic. 

Epigastric hematopoietic nodules were dissected from shrimp as described by 
Martin et al. (1987). Touch preparations of sagittally cut nodules were air-dried prior 
to the cytochemical demonstration of prophenoloxidase (Ppo). Frozen sections (7 
Aim thick) were cut using a Tissue Tek II cryostat for the demonstration of lysosomal 
enzymes. Thin sections (7 /urn) were also prepared using formalin-fixed, par- 
affin-embedded tissue. 

Demonstration of cytoplasmic constituents 

Following a two-minute fixation in absolute ethanol, smears were stained with 
bromphenol blue or periodic acid-Schiff(PAS). Smears to be stained with Best's car- 



1 80 J. E. HOSE ET AL. 

mine (Sheehan and Hrapchak, 1980) were fixed in ethanol for 30 minutes. PAS and 
carmine were compared with and without prior digestion by a-amylase. For the diges- 
tions, hemocytes were suspended in 0.5% aqueous amylase for 1 h, then pelleted by 
a 5 min centrifugation at 500 X g in a table top centrifuge before preparation of the 
smear. Sections of hematopoietic nodule were stained with bromphenol blue or PAS. 

Enzymatic extractions to demonstrate composition of cytoplasmic deposits and 
granules were also examined using EM. Hemocytes were fixed in 2.5% glutaraldehyde 
in 0.1 M sodium cacodylate (pH 7.8) containing 12% glucose for 1 h, then washed in 
cacodylate buffer and kept at 4C for 12 to 18 h. The hemocyte pellet was then dehy- 
drated through a graded ethanol series, infiltrated and embedded in Spurrs' (1969) 
low viscosity plastic. Thin sections were cut on a Porter Blum MT2B ultramicrotome, 
picked up on gold grids, and floated on one of the following solutions for 2 to 20 h at 
37C: (A) 0.5% protease in 0.2 M phosphate buffer (pH 7.4) or (B) 0.5% a-amylase in 
0.2 M phosphate buffer (pH 7.4). These sections and control sections (floated on 
distilled water for an equivalent period of time) were examined unstained and stained 
(0.5% uranyl acetate in 0.05 M Tris-maleate for 1 h at room temperature) using a 
Hitachi HU1 1A transmission electron microscope. 

Lipids were demonstrated in hemocyte smears and nodule touch preparations 
using a commercial Sudan black B kit (Sigma Chemical Co. Kit #380) according to 
provided directions. 

Prophenoloxidase activity 

To test for the presence of prophenoloxidase (Ppo), hemocytes and hematopoietic 
tissue touch preparations were fixed in 2.5%. glutaraldehyde in 0. 1 M phosphate buffer 
(pH 7.4) for 1 h at 4C. The cells were given three 1 5-min rinses in phosphate buffer, 
incubated in 0. 1 % L-DOPA in phosphate buffer for 1 6 h at room temperature (Soder- 
hall and Smith, 1976), and examined by light microscopy. 

Lysosomal enzymes 

The presence of acid phosphatase (Sigma Chemical Co. Kit #386), jfr-glucuroni- 
dase (Kit #180), and a-aryl naphthyl esterase a nonspecific esterase (Kit #90) 
were demonstrated at the light microscopic level using commercial research kits 
(Sigma Chemical Co.). Hemocyte smears and frozen sections of hematopoietic nod- 
ule were fixed in glutaraldehyde and incubated according to provided directions. 
Staining patterns for each enzyme were quantified at 1000X by estimating the num- 
bers and sizes of positive areas in 10 cells from each of the 4 hemocyte categories 
described by Martin el al. ( 1 987). 

To localize acid phosphatase at the EM level, fixed hemocytes and hematopoietic 
nodules were washed thoroughly and then incubated in a medium consisting of 
40 mM Tris-maleate buffer (pH 5), 1 1.5 mM sodium /3-glycerophosphate, 2.4 mM 
lead nitrate, and 5% sucrose at 37C for 2 h. Hemocytes were then processed as de- 
scribed above. 

The same procedure was followed for glucose-6-phosphatase, alkaline phospha- 
tase, and peroxidase except for the use of different incubation media. For glucose-6- 
phosphatase, the fixed cells were incubated in a medium composed of 25 mg glucose- 
6-phosphate, 27 ml distilled water, and 20 ml of 0.3 M Tris-maleate buffer (pH 9.7). 
The incubation medium for alkaline phosphatase consisted of 4 ml 1.25% sodium (3- 
glycerophosphate, 4 ml of 0.2 M Tris-maleate buffer (pH 9), 9.4 ml distilled water, 
and 2.6 ml of 1% lead nitrate. The peroxidase medium contained 5 mg 3,3-diamino- 



CYTOCHEMISTRY OF SHRIMP HEMOCYTES 181 

benzidine tetrahydrochloride, 10 ml of Tris-maleate buffer (pH 7.6), and 0.1 ml of 
1%H 2 O 2 . 

RESULTS 
Cytoplasmic constituents 

The abundant cytoplasmic deposits of agranular hemocytes and a subset of small 
granule hemocytes are composed of glycoproteins as evidenced by positive reactions 
with PAS, carmine, and bromphenol blue. Digestion with a-amylase prior to applica- 
tion of PAS and carmine reduced but did not completely remove the staining of these 
cytoplasmic deposits. Tissue sections of the epigastric hematopoietic nodule stained 
with PAS or bromphenol blue yields results similar to those in free hemocytes. 

Sudan black B produces a distinctive staining pattern in agranular hemocytes and 
small granule hemocytes with cytoplasmic deposits (Figs. 1, 2). These cells appear 
smudgy, with the heavy dark stain obscuring nuclear characteristics. Only a thin clear 
zone adjacent to the plasma membrane is occasionally present. Staining of the cy- 
toplasmic deposits by Sudan black B indicates the presence of a lipid moiety associ- 
ated with the glycoprotein. In small granule hemocytes lacking deposits (Fig. 3) and 
in large granule hemocytes (Fig. 4), delicate staining is evident only around granule 
and nuclear membranes, producing a diffuse pattern. Maturing hemocytes from the 
hematopoietic nodule display identical staining patterns. 

Granule histochemistry 

Granules in free and maturing hemocytes are stained with PAS, carmine, and 
bromphenol blue, indicating the presence of glycoproteins. Prior amylase digestion 
removes granular staining by PAS and carmine. 

Prophenoloxidase activity is visualized following incubations of fixed hemocytes 
and hematopoietic tissue touch preparations with L-DOPA (Figs. 5-8). Ppo activity 
is limited to granules of small granule hemocytes lacking glycoprotein deposits and 
large granule hemocytes. In some animals (molt stage D), almost 100% of these cell 
types display intense activity (> 10 positive granules each) while in intermolt shrimp, 
less than 1% of these cells are positive. Similar results are obtained using hemocyte 
smears and tissue touch preparations. 

No peroxidase activity is observed in any of the hemocyte categories. 

The glycoprotein content of large and small granules is also seen in sectioned 
tissues that were subsequently treated with protease or -amylase. Figure 9 shows a 
large granule from a hemocyte viewed after standard preparation. Figures 10 and 1 1 
show granules in sections treated with a-amylase ( 12 h) and protease (6 h). At these 
times, the core of the granules has been extracted, however, with longer incubations 
(20 h), the entire granule is extracted by both enzymes. 

Lysosomal enzymes 

Three hydrolases (acid phosphatase, /i-glucuronidase, and nonspecific esterase) 
were used to demonstrate the presence of lysosomes in hemocytes at the light micro- 
scope level (Table I). These stains yield similar cytochemical information for each 
specific hemocyte type although individual hydrolases produce slightly different 
staining patterns. Agranular hemocytes do not contain any of the lysosomal enzymes. 
Glycoprotein-rich small granule hemocytes exhibit between zero and three focally 
positive areas consistent with the size of lysosomes. These cells occasionally contain 



182 



J. E. HOSE ET AL. 










10 



11 



FIGURES I -4. Light micrographs of agranular, small granule hemocyte with deposits, small granule 
hemocyte without deposits, and large granule hemocyte, respectively, treated to show sites of prophenoloxi- 
dase. The first two cells have no reaction product. The granules (arrows) in the small granule hemocyte 
without deposits react as does the entire cytoplasm of the large granule hemocyte. All figures 2500X. 

FIGURES 5-8. Light micrographs of same cell types as in Figures I -4, treated with Sudan black B. 
Agranular and small granule hemocytes with deposits show dense reaction products in the cytoplasm which 
obscure the nucleus. The latter two cell types have minimal staining and the nucleus (N) is clearly observed. 
All figures 2500X. 

FIGURE 9. Transmission electron micrograph showing homogeneous and electron-dense granules 
(G) from a large granule hemocyte fixed with both glutaraldehyde and osmium and stained with uranyl 
acetate and lead citrate. 43,OOOX. 

FIGURE 10. Transmission electron micrograph showing a granule from a large granule hemocyte 
that was fixed only with glutaraldehyde. Thin sections were floated on a protease solution for 6 h and 
examined without stain. Note the low electron density and extraction of the granule core (C). 43,OOOX. 

FIGURE 1 1 . Transmission electron micrograph of a granule from a large granule hemocyte prepared 
as in Figure 10 and then floated on a solution of a-amylase for 12 h. Note the low electron density of the 
granule and extraction of its core (C). 43,000x. 



a few acid phosphatase-positive granules as well. In contrast, small granule hemocytes 
lacking cytoplasmic deposits have from three to eight positive foci consistent with 
lysosomes. Half of these cells have only a few (0-3) positive granules while the re- 
maining small hemocytes contain over 30 positive granules. A few of the latter group, 
presumably transitional to large granule hemocytes, also exhibit a few large acid phos- 
phatase-positive granules. In the large granule hemocytes, up to three focally positive 



CYTOCHEMISTRY OF SHRIMP HEMOCYTES 



183 



TABLE I 

Distribution oflysosomal enzymes in shrimp hemocytes 



Hemocyte type 


Acid phosphatase 


/8-glucuronidase 


Glucose-6- Non-specific 
phosphatase esterase 


Alkaline 
phosphatase 


Agranular 


None 


None 


None None 


None 


Small granule 
hemocyte with 
deposits 


Rare(l-3RS*/ 
Cell) 


Rare(l-3RS/ 
Cell) 


None Few(l-10RS/ 
Cell) 


None 


Small granule 
hemocyte without 
deposits 


Mixed (50% of cells 
have>30RS/ 
Cell 50% of cells 


Many 
(>10RS/Cell) 


None I ntermediate ( 1 0- 
30 RS/Cell) 


None 



Large granule 
hemocytes 



have 1-10RS/ 
Cell) 

Mixed (50%. of cells 
have 0-1 RS/Cell 
and nuclei are 
pycnotic 50% of 
cells have 4-8 
RS/Cell) 



Many 
(> 10 RS/Cell) 



None 



Many 
(> 30 RS/Cell) 



None 



* RS stands for reaction sites. 



areas consistent with lysosomes were observed. From zero to five small granules are 
positive as well as from zero to two large granules. Among the large granule hemo- 
cytes, the largest cells which contain eccentrically placed, pycnotic nuclei were usually 
acid phosphatase-negative or contain only one positive focus. 

Electron microscopy localization of acid phosphatase yields similar results with 
no reaction product detected in agranular hemocyte (Fig. 12). Staining is infrequently 
observed in small granule hemocytes containing glycoprotein deposits and is re- 
stricted to small vesicles and granules of the non-striated variety (Fig. 13). Heavy 
staining is found in the granules of the small granule hemocyte lacking deposits (Fig. 
14). Large granule hemocytes have reaction product dispersed throughout the cell in 
vesicles and the smaller granules. Only a few of the large granules stain positive al- 
though these were morphologically indistinguishable from non-reactive granules 
(Fig. 15). In Figure 16, acid phosphatase-positive trans cisternae and small vesicles 
are shown budding from a Golgi body. These vesicles (Fig. 16, inset) appear to pro- 
gressively coalesce, forming larger reaction vesicles (Fig. 1 7), then small granules, and 
finally large granules (Fig. 1 8). 

Nonspecific esterase is observed only in granulated cells. Glycoprotein-rich small 
granule hemocytes are completely negative or contain up to 10 tiny positive areas 
consistent with the size of vesicles. In contrast, from 10 to over 30 positive vesicles 
are observed in small granule hemocytes without cytoplasmic deposits. Large granule 
hemocytes contain numerous (>30) positive vesicles. Patterns of /3-glucuronidase 
staining in granulated cells are similar to those of nonspecific esterase in addition 
to the presence of a few (<3) positive foci of lysosomal size in the small granule 
hemocytes. 

Maturing hemocytes from frozen sections of the hematopoietic nodule were ex- 
amined for the presence of acid phosphatase and /3-glucuronidase. Staining patterns 
of acid phosphatase are identical between maturing hemocytes and those described 
above for free hemocytes. 0-glucuronidase activity is not observed in touch prepara- 
tions of the hematopoietic nodule. 

All hemocytes are negative for alkaline phosphatase and glucose-6-phosphatase. 



184 



J. E. HOSE ET AL. 














15 








G 



16 



17 



18 



CYTOCHEMISTRY OF SHRIMP HEMOCYTES 185 

DISCUSSION 

Results of cytochemical tests support the morphological classification of ridge- 
back prawn hemocytes previously developed in our laboratory (Martin et al., 1987) 
and yield information on the physiological functions performed by the various hemo- 
cyte types. A combination of two or three cytochemical tests is suggested for classifi- 
cation of shrimp hemocytes. Sudan black B produces a distinctive smudgy staining 
pattern in agranular hemocytes and small granule hemocytes with cytoplasmic de- 
posits. Acid phosphatase can be used to differentiate agranular cells, which are nega- 
tive for lysosomal enzymes. Prophenoloxidase activity is limited to small granule 
hemocytes without cytoplasmic deposits and large granule hemocytes; however, sig- 
nificant activity may only be demonstrable during the D stage of the molt cycle (Bau- 
chau, 1981). 

The glycoprotein deposits in the cytoplasm of agranular hemocytes and a sub- 
group of small granule hemocytes are distributed in linear arrays throughout the en- 
tire cell and are evident in all molt stages (unpub. obs.). In contrast, glycogen which 
it resembles ultrastructurally is typically confined to one area of decapod hemocytes 
and does not have a linear arrangement (Johnston et al., 1973; Bauchau, 1981). Gly- 
cogen has been shown to be transported by hemocytes (Johnston et al., 1973; Bau- 
chau, 1981) and may involve the enzyme glucose-6-phosphatase (Johnston and Da- 
vies, 1 972). This enzyme, however, was not detected in shrimp hemocytes. The glyco- 
protein may contain a lipid moiety since the deposits are intensely stained by Sudan 
black B. Such chemical properties are consistent with those of the primary coagula- 
tion protein, coagulogen (Durliat, 1985). In decapod hemocytes, intracellular coagu- 
logen does not appear to be localized in granules although granules are necessary 
for coagulation to occur (Ravindranath, 1980; Durliat, 1985). Shrimp small granule 
hemocytes with lipoglycoprotein deposits contain granules with a striated or concen- 
tric substructure (Martin et al., 1987). Similar granules have been observed in Limu- 
lus (Copeland and Levin, 1 985), crabs (Bodammer, 1 978), lobsters (Hearing and Ver- 
nick, 1967; Goldenberg et al., 1986), and crayfish (Unestam and Nylund, 1972), and 
alterations in the striated granules of shrimp have been observed early in the process 
of hemolymph coagulation (unpub. obs.). 

Lysosomes were observed in each cell type except for agranular hemocytes. Small 



FIGURES 12 AND 13. Transmission electron micrographs of an agranular hemocyte (Fig. 12) and a 
small granule hemocyte with deposits (Fig. 13) treated to display sites of acid phosphatase activity. No 
reaction sites are present in agranular cells. In small granule hemocytes with deposits, reaction sites (X) are 
rare and then localized to granules of the electron-dense variety. Striated granules (S) are never labelled. In 
both cells, note the small amount of cytoplasm which contains the deposits (arrows). Both figures 20.000X. 

FIGURE 1 4. Transmission electron micrograph of a small granule hemocyte that lacks deposits show- 
ing a few reaction sites for acid phosphatase in granules (G) and vesicles (V). 20.000X. Inset shows a higher 
magnification (43,OOOX) micrograph of the small granules. 

FIGURE 15. Transmission electron micrograph of a large granule hemocyte showing reaction sites 
for acid phosphatase in vesicles throughout the cytoplasm (arrows) and some of the granules (G). Other 
granules (X) show no reaction product. 20.000X. 

FIGURE 16. Transmission electron micrograph showing a Golgi body in a large granule hemocyte. 
The trans-cisternae contains the acid phosphatase reaction product. 42,000x. Inset shows a vesicle with 
reaction product. Similar vesicles are commonly seen around Golgi bodies as well as throughout the cyto- 
plasm. 42.000X. 

FIGURE 1 7. Transmission electron micrograph showing acid phosphatase reaction product in three 
vesicles of increasing diameter. Note how the smallest vesicle appears to be fusing (arrow) with the medium 
sized vesicle and that the contents of the vesicles are not as electron dense as fully mature granules. 32,000x. 

FIGURE 18. Transmission electron micrograph showing acid phosphatase reaction product in one 
large vesicle (V) and not in two adjacent granules (G). 32,OOOX. 



186 J- E. HOSE ET AL. 

granule hemocytes with glycoprotein deposits contained only one to three lysosomes 
per cell. In small granule hemocytes without deposits and in large granule hemocytes, 
many lysosome e identified using LM and EM cytochemistry. Although the gran- 
ules in thes s are morphologically indistinguishable, they may be enzymatically 
heteroge s (see Bauchau, 1981). Using TEM, acid phosphatase was localized in 
some iot all of the granules of the shrimp. The same results were observed in 
5 of the clam Mercenaria mercenaria (Yoshino and Cheng, 1976) and 
interpreted to indicate a heterogeneous population of granules or a non-synchronized 
cycle of granule production, perhaps with the final enzyme stored in an inactive form. 
The presence of numerous lysosomes in the large and small granulocytes which lack 
deposits supports the suggestion that these cells are phagocytic (Bauchau, 1981; Sod- 
erhall et al, 1986) and that the granules are available for intracellular degradation 
processes. However, because of the large number of granules in a single hemocyte, it 
is unlikely that granules could be exclusively reserved for phagocytosis. Other re- 
searchers suggested extracellular functions for these granules, including recognition 
of foreign material (Soderhall and Smith, 1983) and agglutinin sequestration (Stang- 
Voss, 1971). 

The recognition of foreign material in arthropods is mediated by the propheno- 
loxidase system which is located in the granules (Soderhall and Smith, 1983). Results 
of the present study show that only large granule hemocytes and small granule hemo- 
cytes without deposits contain prophenoloxidase activity. Soderhall and Smith 
( 1 983) obtained similar conclusions of Ppo activity within granular hemocytes in the 
crab, Carcinus maenus. Ppo activation and exocytosis in response to endotoxin or @- 
glucan exposure can initiate the coagulation cascade and serves as the crustacean 
equivalent to the alternate complement and properdin pathways in mammals (Dur- 
liat, 1985). The proposed existence of the Ppo system in the small granule hemocytes 
without deposits-large granule hemocyte line and the coagulation enzymes in the 
other hemocyte line suggests cooperativity among shrimp hemocytes during endo- 
toxin-mediated coagulation analogous to that observed for defense reactions in in- 
sects (Ratcliffe et al., 1984) and crustaceans (Soderhall et al., 1986). 

Based upon results of morphological (Martin et al., 1987) and cytochemical stud- 
ies, shrimp hemocytes can be divided into two cell lines, the deposit line (composed 
of agranular and striated granule hemocytes) and the granulocyte line (small and 
large granule hemocytes). Although the lysosomal enzyme data presented here are 
consistent with maturing stages of a single cell line, the following arguments support 
our theory: ( 1 ) striated granules are never found in granulocytes (The striated granule 
shown in Fig. 7B of Martin and Graves, 1985, was taken from a small granule hemo- 
cyte containing cytoplasmic deposits. However, at that time, no distinction was 
made between the deposit and granulocyte lines.); (2) glycolipoprotein deposits are 
never observed in granulocytes; (3) mitosis is observed both in agranular hemocytes 
and in small granule hemocytes which lack cytoplasmic deposits; (4) cells are present 
as a continuum of differentiation between agranular and striated granule hemocytes 
and between small and large granule hemocytes; and (5) clusters of deposit cells and 
granulocytes are usually segregated within the hematopoietic tissue. The utility of this 
classification scheme must now be determined by functional studies identifying the 
role of the various hemocyte types in crucial biological processes such as coagulation, 
defense reactions, wound healing, and exoskeleton hardening. 

ACKNOWLEDGMENTS 

We thank Terri Fu and Sidne Omori for their technical help. The project was 
supported by NSF grant DCB-8502 1 50 to GM and JEH. 



CYTOCHEMISTRY OF SHRIMP HEMOCYTES 187 

LITERATURE CITED 

ANDERSON, S. L. 1985. Multiple spawning and molt synchrony in a free spawning shrimp (Sicvonia in- 

ge/rtw.-Penaeoidea). Biol. Bull. 168: 377-394. 
BAUCHAU, A. G. 1981. Crustaceans. Pp 386-420 in Invertebrate Blood Cells, Vol. 2. Academic Press, New 

York. 
BODAMMER, J. E. 1978. Cytological observations on the blood and hemopoietic tissue in the crab, Calli- 

nectes sapidus. I. The fine structure of hemocytes from intermoult animals. Cell. Tissue Res. 187: 

79-86. 
COPELAND, D. E., AND J. LEVIN. 1985. The fine structure of the amoebocyte in the blood of Limitlus 

polyphemus. I. Morphology of the normal cell. Biol. Bull. 169: 449-457. 
DURLIAT, M. 1985. Clotting processes in Crustacea Decapoda. Biol. Rev. 60: 473-498. 
GOLDENBERG, P. Z., A. H. GREENBERG, AND J. M. GERRARD. 1986. Activation of lobster hemocytes: 

cytoarchitectural aspects. J. Invertebr. Pathol. 47: 143-154. 
HEARING, V. J., AND S. H. VERNICK. 1967. Fine structure of the blood cells of the lobster, Homarus 

americanus. Ches. Sci. 8: 1 70. 
JOHNSTON, M. A., AND P. S. DAVIES. 1972. Carbohydrates of the hepatopancreas and blood tissues of 

Carcinus. Comp. Biochem. Physio/. 41 B: 433-443. 
JOHNSTON, M. A., H. Y. ELDER, AND P. S. DAVIES. 1983. Cytology of Carcinus haemocytes and their 

function in carbohydrate metabolism. Comp. Biochem. Physiol. 46A: 569-581. 
MARTIN, G. G., AND B. L. GRAVES. 1985. Fine structure and classification of shrimp hemocytes. /. Mor- 

phol. 185: 339-348. 
MARTIN, G. G., J. E. HOSE, AND J. J. KIM. 1987. Structure of hematopoietic nodules in the ridgeback 

prawn, Sicvonia ingentis: light and electron microscopic observations. J. Morphol. 192: 193-204. 
RATCLIFFE, N. A., C. LEONARD, AND A. F. ROWLEY. 1984. Prophenoloxidase activation: nonself recogni- 
tion and cell cooperation in insect immunity. Science 226: 557-559. 
RAVINDRANATH, M.H.I 980. Haemocytes in haemolymph coagulation of arthropods. Biol. Rev. 55: 1 39- 

170. 
SHEEHAN, D. C., AND B. B. HRAPCHAK. 1 980. Theory and Practice ofHistotechnology. C. V. Mosby Co., 

St. Louis. 48 1 pp. 
SODERHALL, K., AND V. J. SMITH. 1983. Separation of haemocyte populations of Carcinus maenus and 

other marine decapods, and prophenoloxidase distribution. Dev. Comp. Immunol. 7: 229-239. 
SODERHALL, K., V. J. SMITH, AND M. W. JOHANSSON. 1986. Exocytosis and uptake of bacteria by isolated 

haemocyte populations of two crustaceas: evidence for cellular co-operation in the defence reac- 
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SPURRS, A. 1969. A low viscosity epoxy embedding medium for electron microscopy. J. Ultrastrucl. Res. 

26:31-43. 
STANG-Voss, C. 1971. Zur ultrastruktur der blutzellen wirbelloser tiere. V. Liber die hamocyten von Asta- 

cus astacus (L.) (Crustacea). Z. Zellforsch. 122:68-75. 
UNESTAM, T., AND J.-E. NYLUND. 1972. Blood reactions in vitro in crayfish against a fungal parasite, 

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Reference: Biol. Bull. 173: 188-204. (August, 1987) 



IMPULSE PROPAGATION AND CONTRACTION IN THE TUNIC 

OF A COMPOUND ASCIDIAN 

G. O. MACKIE AND C. L. SINGLA 

Department of Biology, University of Victoria, Victoria. British Columbia, Canada V8W 2Y2 

ABSTRACT 

Diplosoma listerianum and D. macdonaldi (Earn. Didemnidae) have a network 
of cells ("monocytes") in the tunic which contain high concentrations of microfila- 
ments and react positively with NBD-phallacidin, indicating the presence of F-actin. 
The tunic is contractile, especially in the areas around the cloacal apertures, which 
can be closed completely. Myocytes are concentrated in sphincter-like bundles 
around these openings, but also are found throughout the tunic. Electrophysiological 
recordings reveal a diffuse conduction system in the tunic propagating all-or-none 
impulses ("tunic potentials," TPs) through all parts with a conduction velocity of 
< 1 .5 cm s ', and a refractory period of 1 .6 s. TPs correlate one-for-one with contrac- 
tions. The system is excitable to the touch, but is also spontaneously active, showing 
steady patterns of potentials as well as regular, 'parabolic 1 bursts. The evidence sug- 
gests that the myocyte net itself conducts the impulses triggering the contractions. In 
the absence of conventional nerves and muscles, the system provides the colony with 
a way of regulating the effluent water current and hence the volume of a common 
cloacal space. 

The TP system is not 'wired in' to the ascidiozooids either as a sensory or as a 
motor pathway. The tunic acts as an independent behavioral entity. 

INTRODUCTION 

The ascidian tunic or test is "an outer covering which completely surrounds the 
individual zooid in solitary ascidians or forms a common groundwork in which the 
zooids are embedded in colonial species." (Goodbody, 1974). It is a secretion product 
of the body wall epithelium and consists of a matrix of proteins and carbohydrates 
(including cellulose) into which cells migrate from the hemocoel during develop- 
ment. Blood vessels often penetrate the tunic, and sensory processes from receptors 
whose cell bodies lie in the underlying epithelium may also extend into the tunic 
(references in Bone and Mackie, 1982) but muscles and nerves' are absent. The vari- 
ous cells present may be concerned with secretion of tunic materials, phagocytosis, 
self-nonself discrimination, coloration, and some other less well understood func- 
tions. Some tunic cells are capable of movement and have contractile pseudopodia or 
filopodia, but the contractions reported are very slow (< 1 14 ^m per hour in Botryllus 
according to Izzard, 1974). Several authors (e.g., Saint-Hilaire, 1931;Godeaux, 1964) 
have likened the tunic to mesenchyme. Brien ( 1 966) calls it "a living envelope, equiv- 
alent to a sort of peripheral mesenchyme." Unlike mesenchyme, however, it is not 



Received 9 April 1987; accepted 26 May 1987. 

' There appears to be only one report of nerve cells in the tunic of an ascidian, that of Das (1936). No 
later study on tunic histology supports this claim. 

188 



TUNIC RESPONSE SYSTEM 189 

covered by epithelium but is exposed to the environment, and in this respect it more 
resembles a cuticular or exoskeletal tissue. 

Given the absence of nerves and muscles from the tunic, it is not surprising that 
there have been no reports that the structure responds to stimulation, contracts, or 
'behaves' in the usual sense, although in several cases it is composed of a fairly plastic, 
viscous material capable of short-term conformational changes (Delia Valle, 1908; 
Godeaux, 1964). During observations on Diplosoma listerianum, however, it became 
clear that this species has a tunic in which electrical signals propagate on an all-or- 
nothing basis, mediating contractions of the tunic itself. In this report, the electro- 
physiological characteristics of this conduction system are described, along with an 
account of the activities performed and of the cells likely responsible for conduction 
and contraction. The evidence implicates a novel type of cell ("rnyocyte") as the basis 
for both conduction and contraction. These cells seem to combine the properties of 
conventional nerves and muscles including the ability to function as pacemakers. 
They are distributed throughout the whole tunic in the form of a dense network 
which, it is proposed, constitutes the structural basis for the behavioral action system 
whose electrical correlates are picked up with recording electrodes. 

MATERIALS AND METHODS 

Two species of Diplosoma were used in this study. D. listerianum Milne-Edwards, 
1841, was obtained at the Stazione Zoologica in Naples, Italy. A species tentatively 
identified as D. macdonaldi Herman, 1 886 was obtained at the Friday Harbor Labo- 
ratories of the University of Washington, and at the Bamfield Marine Station, Barn- 
field, British Columbia, Canada. D. macdonaldi and D. listerianum are very similar 
and may be conspecific (Monniot, 1 974). The specimens collected at Naples grew on 
the walls of the public display aquarium and elsewhere in the seawater system, where 
they appear to be endemic. D. macdonaldi specimens were collected from rocks and 
pilings in the intertidal zone. Following the method of Delia Valle ( 1 908 ), specimens 
were removed from their natural substrates and transferred to glass slides or petri 
dishes. There they attached after a few hours, subsequently resumed growth, ex- 
panded and put out new attachment structures ("crampons" 1 ). All the experiments 
reported in this paper were performed on transplanted specimens maintained in run- 
ning seawater in the laboratory. The bulk of the work was done at Naples, and 
D. listerianum was used for all the illustrations except Figures 2, 8, and 9. 

For histological study, pieces of tunic were dissected out and mounted as whole 
mounts either fresh or after fixation and examined by phase contrast or Nomarski 
differential interference contrast microscopy. NBD-phallacidin (from Molecular 
Probes Inc., 24750 Lawrence Road, Junction City, Oregon 97448) was used as a 
specific fluorescent stain for F-actin. Material was embedded in Epon 8 1 2 for electron 
microscopy after standard fixation and processing. 

Electrophysiological study was carried out on small, whole colonies which had 
become well established on their glass or plastic substrates. A slow flow of water was 
maintained through the preparation dish during the experiments to ensure that the 
colonies behaved as nearly as possible as in nature. Thus, temperatures in the prepara- 
tion dish were kept close to those in the seawater systems at the laboratories where the 
animals were maintained ( 17-19C at Naples, 1 1-1 3C at Friday Harbor). A simple 
thermistor flow meter (Mackie et ai, 1983) was used to record changes in water flow 
velocity out of the cloacal apertures. For stimulation and recording, polyethylene 
suction electrodes were used. Signals were amplified and displayed on an oscilloscope 
or on a chart recorder. For consistency with our earlier papers on tunicate electro- 



190 



G. O. MACKIE AND C. L. SINGLA 



cloacal aperture 



zooid 

common cloaca 




crampon " 

FIGURE 1 . Diplosoma listerianium, cut-away drawing after Lahille ( 1 890). The zooids hang by their 
oral siphons from the upper tunic layer and are anchored below by strands of tunic drawn up from the 
basal tunic layer, which is attached to the substrate by "crampons." Arrows show water flow. 



physiology (Bone and Mackie, 1982) the polarity of the electrical records is arranged 
so that negative events go up, positive down. 

General description of Diplosoma and its activities 

In Diplosoma and other didemnids the tunic is drawn out into thin sheets an 
upper sheet from which the zooids are suspended and a lower (basal) sheet which 
attaches to the substrate (Fig. 1 ). The tunic material composing these sheets is directly 
exposed to the seawater on both sides, and lacks an epithelial covering. A thin layer 
of tunic encases the zooids (depicted by Carlisle, 1953) and this continues down into 
an attachment strand ("stalk") which anchors the zooid to the basal tunic sheet. A 
retractor muscle and fine blood vessels (30 /^m diam.) pass down the stalk from the 
zooid. It is incorrect to refer to the stalk as the retractor muscle (e.g., Berrill, 1950) as 
it is composed primarily of tunic, and the muscle penetrates it for only a short dis- 
tance. The blood vessels entering the stalk, typically four (Pizon, 1905), enter the 
basal sheet and run out into it, terminating in vascular ampullae. The ampullae con- 
tract and expand, pulsating rhythmically as in other ascidians, but never swell to 
more than 250 ^m in diameter. Contrary to the arrangement in colonial styelids such 
as Botryllus, the blood vessels of different zooids are not interconnected. The vascular 
ampullae are responsible for the formation of 'crampons' (ramponi, Wurzeln}: spe- 
cialized patches of tunic material 180-240 ^m in diameter by which the basal tunic 
adheres to the substrate. The ampullae, along with their blood vessels, may withdraw 
after the crampons are complete, leaving behind an attachment strand of pure tunic 
material. These strands are most conspicuous around the edges of the colony (Fig. 
2). When elongated, they resemble the guy-ropes of a tent (Carlisle, 1 96 1 ). Crampons 
are also present underneath the colony, roughly four per zooid stalk. 

Water enters the colony through the oral siphons of the ascidiozooids. As the 
zooids lack atrial siphons, water passes directly out into the common cloacal cavity 
from which it finally exits via large cloacal apertures, which are often more than 1 
mm in diameter. The exhalent water forms a plume that may rise to a considerable 
height above the surface of the colony. Small apertures (<150 ^m) are also present 
in the basal tunic (Fig. 1) and water passes through them into the narrow space be- 
tween the tunic and the substrate and then to the exterior. The cloacal apertures are 
simply holes in the tunic and should not be referred to as siphons, as they are not 
parts of zooids. A single large cloacal aperture may serve as the exhalent water route 



TUNIC RESPONSE SYSTEM 



191 




FIGURE 2. Diplosoma macdonaldi. A. Portion of a colony seen from above, showing a cloacal aper- 
ture (ca), crampons (cr), and zooids, some with their oral siphons (os) in focus. B. Enlargement of edge, 
showing two crampons, both containing vascular ampullae. The one on the left (am) is expanded, while 
the one on the right which comes from another zooid is contracted. Arrows show the blood vessel of 
the ampulla on the right. 



for some 50 zooids. Stimulation of the tunic at any point results in slow closure of 
the cloacal apertures, a response discussed in detail below. 

A well-maintained colony which is actively feeding and growing in undisturbed 
conditions tends to be flat, the stalks of its zooids very short (<100 /urn), and the 
common cloacal space relatively small. The blood vessels passing down the zooid 
stalks extend well out into the basal tunic. Around the edges of the colony these 
vessels push out and form crampons (Fig. 2B). In colonies which are not feeding and 
growing so vigorously or which have been kept in stagnant water for a few hours, the 
blood vessels retract and retreat up the stalks into their zooids. trailing their ampullae 
behind them. At the same time, the stalks elongate and are drawn out into thin 
strands 1 mm or more in length. Elongation of the stalks accompanies swelling of the 
cloacal space with exhalent water, and the whole colony expands. These changes, 
documented in part by Delia Valle (1908, and earlier papers cited), seem to be a 
response to changed water conditions, but it is interesting to learn that in Diplosoma 
virens expansion and contraction are periodic events exhibited according to a diurnal 
rhythm (J. S. Ryland, pers. comm.). 

Didemnid colonies are known to be capable of locomotion (e.g., Delia Valle, 
1908; Carlisle, 1961; Ryland et al, 1984). The exact mechanism of locomotion has 
never been properly analyzed, but it involves the projection of finger-like tunic pro- 
cesses containing blood vessels, whose ampullae form new crampons at attachment 
sites ahead of the colony in the direction of movement. At the rear end of the moving 
colony these attachment processes, vacated by their blood vessels, are stretched out 
thin and eventually detach or break off. There is some evidence of positive phototaxis: 
Delia Valle (1908) found that colonies tended to move upward in the public display 
tanks at Naples which are lit from above stopping only when they reached the 
surface. Carlisle (1961) found that Diplosoma moved sideways when illuminated 
from the side. Crampons once formed cannot be lifted up and moved to another site, 
so the movement cannot be thought of as a type of 'walking'; rather, it resembles the 



192 G. O. MACKIE AND C. L. SINGLA 

motion of a tracked vehicle, a slow flowing over fixed points which presumably re- 
quires secretion of new tunic at the advancing end. The process requires further study. 
The asckUozooids ofDiplosoma behave like solitary ascidians (reviewed by Bone 
and Mackie, 1982), pumping water continuously when not disturbed, and contract- 
ing their oral siphons and arresting their cilia in response to mechanical interference, 
as for instance with the entry of an excessively large food particle. Stronger mechani- 
cal stirnuSation causes retraction of the whole zooid by the retractor muscle which 
runs down into the proximal part of the stalk. These activities are carried on indepen- 
dently by the zooids. Stimulation does not cause the spread of zooid contractions or 
ciliary arrests across the colony. This is in marked contrast to the situation in Bortryl- 
lus and its relatives, where signals propagate through the colonial network of blood 
vessels triggering behavioral events in the zooids (Mackie and Singla, 1983). 

Histology 

The living tunic is soft, pliable, and transparent. The ground substance shows no 
regional differentiation except at the surfaces, where there is a thin (50 nm) cuticular 
layer comparable to the "outermost cuticle" ofdona tadpole larvae (Gianguzza and 
Dolcemascolo, 1984), but bearing a fuzzy surface coating 200 nm thick. There ap- 
pears to be no counterpart to the subcuticular zone seen in adults of this and other 
solitary ascidian species (De Leo et ai, 1981; D'Ancona Lunetta, 1983), but a layer 
about 200 nm deep underlying the cuticle is more densely fibrous than in other re- 
gions. Calcareous spicules are present (Carlisle, 1953) but are extremely small (<10 
^m) and far apart. Conspicuous in all parts of the tunic are the large, spherical, vacuo- 
lated cells termed "kalymmocytes" by Salensky ( 1 892) which are probably the coun- 
terparts of the bladder cells (Blasenzelleri) or Saint-Hilaire (1931) and the cellules 
vesiculeuses of Godeaux (1964). Peres (1948) one of the few authors to study post- 
larval Diplosoma calls them "lacunae," which is clearly inappropriate, as they are 
cells, not spaces. Also present are cells resembling the granulocytes, morula cells, 
phagocytes, and other immigrant blood cells described in various tunicates by various 
authors. Much uncertainty surrounds the identification of such cells, but this is irrele- 
vant to the present discussion. Bacteria are usually present in the tunic ground sub- 
stance. 

Of particular interest in the context of the present investigation are two cell types, 
both with processes interconnecting to form networks. Neither of these is clearly iden- 
tifiable on the basis of previous descriptions, so they will be given new names: filo- 
podial cells and myocytes. Filopodial cells (Fig. 3A) are restricted to the surface layer 
of the tunic, while the myocytes lie deeper. Filopodial cells are flattened in the plane 
of the surface layer, with three or more broad cytoplasmic expansions resembling 
neuronal growth cones, each of which subdivides into numerous fine filopodia. The 
filopodial cells form a fairly regular network, and are spaced out so that the filopodia 
just make contact. The cells termed myocytes (Fig. 3B, C) are usually bi-, tri- or 
multipolar, with thicker, much longer processes than the filopodial cells. Their pro- 
cesses show few branches, and rarely subdivide to form filopodia. They are fairly 
straight, and run for considerable distances through the territories of adjacent myo- 
cytes, making numerous contacts with other such processes. The myocyte layer is 
thick, not two-dimensional like that of the filopodial cells. The myocytes are present 
in all parts of the tunic but are concentrated into sphincter-like bundles around the 
cloacal apertures (Fig. 3C) and around the necks of the ascidiozooids. Their presence 
and circular orientation in these places strongly implicates them in the role of the 
contractile elements responsible for constricting the cloacal apertures and for pulling 



TUNIC RESPONSE SYSTEM 193 

in the tunic over the ascidiozooids when retracted, hence the designation "myocyte." 
The filopodial cells seem less likely to fulfill such a role, as they show no such concen- 
trations around the openings, and because their processes seem too delicate to be 
effective as contractile elements. 

Material stained with NBD-phallacidin and examined under a fluorescence mi- 
croscope at 460 nm showed the myocytes as uniformly fluorescent objects, indicating 
the presence of F-actin (Fig. 4A). Kalymmocytes also reacted positively, but other 
cells in the tunic showed little response. The filopodial cells showed a very weak fluo- 
rescence, and only their thicker processes could be seen at all. 

Under the electron microscope (Fig. 5), the myocytes are characterized by dense 
masses of rather loosely arranged fine microfilaments. True smooth muscle in ascidi- 
ans by contrast shows thick and thin myofilaments arranged in strictly parallel arrays 
(Nevitt and Gilly, 1 986). Further, using NBD-phallacidin, true muscle from the man- 
tles of the ascidiozooids in Diplosoma showed a much stronger fluorescent reaction 
(Fig. 4B) than was apparent in myocytes in the same preparations. For these reasons, 
and because of their arrangement in the form of a diffuse plexus, it seems appropriate 
to recognize the myocytes as a new cell type distinct from conventional smooth 
muscle. 

As noted, the filopodial cells and the myocytes lie in different layers of the tunic, 
and show few points of contact; therefore, while it is conceivable that the filopodial 
cells could represent a primitive, neuroid conduction network mediating responses 
of the myocytes, the likelihood of this seems remote. However, we do not know what 
function the filopodial cells serve. 

RESULTS 
The tunic pulse system: basic properties 

Colonies growing in good condition on slides sometimes show no electrical activ- 
ity in the tunic. Usually, however, it is possible to detect spontaneous patterns of 
small electrical impulses (tunic pulses, TPs) which propagate without decrement 
through all parts of the tunic, showing no alteration in wave form even when recorded 
at distances of several centimeters from the site of initiation. They can be conducted 
through narrow bridges of tunic less than 0.5 mm wide. They are exhibited in newly 
settled colonies with only two or four zooids. TPs may be evoked by tactile and elec- 
trical stimulation of the tunic as well as appearing spontaneously. Their characteris- 
tics may be summarized as follows: 

Wave form. When the electrode is first attached it may be impossible to detect 
any signal above the noise level, as a dense plug of tunic must first fill the tip of the 
electrode. However, once the electrode is well attached, and usually after 30 minutes, 
signals can be recorded without difficulty for hours or even days. With fine suction 
electrodes (ca., 50 yum I.D.) attached to the surface of the tunic, the signals are re- 
corded as initially positive-going events rarely exceeding 100 v\ in amplitude, with 
a small but long-lasting negative after-potential (Fig. 6A), and a total duration of 
about 2.5s. With larger-bore electrodes the events are more symmetrically biphasic. 
Recordings from the inner and outer surfaces show similar TP wave forms and ampli- 
tudes. Attempts to record intracellularly from the myocytes failed, so interpretation 
of these extracellularly recorded events is difficult, but they are probably compound 
action potentials representing the summed depolarizations of many conducting ele- 
ments. Somehow, the topography of the electrode attachment area converts these 
summed negative events into a predominantly positive-going signal. With a fine elec- 



194 



G. O. MACKJE AND C. L. SINGLA 







r - , 



E 

o 



f 

OQ 



C 
D 



<u 



<u 

O 



ts> 



<u 
C 



c 

<Sl 



S 




o 



S a 



^ n, 

o ^ 

"3 

o 






r 



O 

CNJ 



t/3 ^ 

O \- 

1! 

U.S 







c 

rn <- 

si 



u 

aj 
O 



TUNIC RESPONSE SYSTEM 



195 




FIGURE 4. NBD-phallacidin: A. Fluorescent reaction in myocyte net (arrowheads), and in shrunken 
kalymmocytes (k); B. In conventional muscle from mantle wall of a zooid. 



trode there would be relatively few conducting elements contributing to the signal, 
and they would tend to fire in synchrony so the wave form shown in Figure 6A may 
closely approximate the fundamental event recorded d.c. from a single cell. 

Slow conduction. Conduction velocity is 1.0-1.5 cm-s ' at 19C (Fig. 6B). No 



. 




0-2 (Jm 




2 pm 



FIGURE 5. Electron micrographs of a myocyte (A) and its process enlarged (B), showing fibrillar 
contents. Bacteria (b) are often present in the tunic ground substance. 



196 G. O. MACKIE AND C. L. SINGLA 




J 





FIGURE 6. Tunic pulses (TPs). A. Spontaneous TP recorded under optimal conditions with a fine 
(50 Mm I.D.) extracellular suction electrode (scale bars: 1 s, 100 /*V). B. A TP recorded sequentially from 
the inside of the basal sheet of the tunic (upper trace) and from the outside of the upper sheet (lower trace) 
following a shock (*) on the basal sheet. Recording electrodes were 3 mm apart, conduction velocity 1.3 
cm-s ' (scale bars: 100ms, 50 ^V). C. With two shocks (*) 1.6s apart, a response was elicited only to the first 
shock (upper trace). When the interval between shocks was increased to 1.8 s, both shocks were followed by 
TPs (scale bars: 0.5 s, 50 ^V). D. A mechanical stimulus (arrowhead) elicited a burst of TPs (scale bars: 10 
s, 200 j/V). 



significant variations in conduction velocity were observed in different parts of the 
tunic. Conduction time increases markedly with successive shocks. With shocks at 7 
s intervals, conduction time increased by 50% of its initial value after only 6 pulses. 
It is not clear if increasing conduction time is due to slower conduction or to passage 
of impulses via less direct routes. 

Long refractory period. At 19C, the absolute refractory period was 1 .6 s (Fig. 6C). 
In the figure, a second response was obtained with two shocks 1.8 s apart, but the 
amplitude of the second TP was considerably reduced, and showed a longer latency. 
A long refractory period would be expected if the action potential has a long duration, 
as proposed above. 

Mechanical and electrical excitability. TPs can be evoked by pinching or pricking 
the tunic (Fig. 6D) or by delivering electrical shocks through a suction electrode at- 
tached to it. As with the recording electrodes, a plug of tissue must fill the tip of the 
stimulating electrode firmly before experiments can begin. Large shocks are needed, 
undoubtedly due to current shunting through the aqueous component of the tunic. 
Responses can usually be obtained with stimulator settings of 30-50 V, 2-5 ms, using 
an electrode with an internal tip diameter of about 120 /^m. Chemical sensitivity was 
not examined in detail, but the mucus and body fluids of a small keyhole limpet 
(species undetermined) which is the most obvious predator ofDiplosoma colonies in 
the seawater system at the Stazione Zoologica had no effect on spontaneous pulse 
patterns recorded from the tunic. The TP system does not appear to be affected by 
changes in light intensity, but this aspect also needs further study. 

Spontaneity. Specimens studied in as near to natural conditions as attachment of 
electrodes would allow showed either (a) absence of all electrical activity, (b) steady, 
almost metronomic pulse trains going on for periods of hours in some cases, typically 
with TPs 7-10 seconds apart (Fig. 7 A), or (c) bursts of TPs repeated at regular inter- 



TUNIC RESPONSE SYSTEM 



197 



t 







1 










' 


J 




i 






1 


! 




I 




| 





B 

FIGURE 7. Spontaneous TP patterns. A. Steady pulse pattern (scale: 10 s). B. 'Parabolic' burst (scale: 
5 min). C. Resetting of steady TP pattern by delivering shocks (*) to produce premature firing of the system. 
Note that the TP elicited by the shock and the one following are both of reduced amplitude (scale: 1 min.) 
D. Termination of steady, spontaneous TP pattern by electrical stimulation (*), causing a high frequency, 
artificial TP burst (6 TPs, 6 seconds apart, scale: 30 s). 



vals. These sequences vary considerably, but in typical long-term burst patterns, the 
bursts last about 10-18 minutes (Fig. 7B), comprise 20-35 individual TPs, and are 
followed by 10-18 minutes of silence before the next burst. Spike frequency increases 
during the early part of the burst and decreases toward the end. If spike frequencies 
are plotted graphically, the curve approximates to a parabola. Parabolic bursting is 
typical of many pacemakers e.g., many molluscan neurons (Strumwasser, 1968). In 
a preparation exhibiting a steady TP pattern, delivering a shock slightly before the 
next predicted spontaneous event resets the pacemaker (Fig. 7C). A steady TP rhythm 
can be terminated or interrupted by stimulating the preparation at a frequency greater 
than the rhythm frequency (Fig. 7D). 

The ability to produce pulse trains and burst patterns is not restricted to any par- 
ticular part of the tunic. Small pieces of tunic with no zooids in them from various 
parts of the upper and basal sheets produced rhythms similar to those seen in intact 
colonies. 

Effect of elevated Mg 2+ . TP rhythms continued unaffected in 81 mM Mg 2+ . In 
105 rrLMMg 2+ , spontaneous TP patterns ceased, but the system could still be excited 
electrically. In 1 50 mMMg 2+ , all TP activity ceased. These findings suggest that either 
conduction, contraction, or junctional transmission in the myocyte network is de- 
pendent on extracellular calcium, as magnesium ions block calcium channels (Hagi- 
wara and Takahashi, 1 967). 

Effect of curare and acetylcholine. Tubocurarine chloride had no effect on the 
wave forms of TPs nor on their spontaneous patterns when used at concentrations 
up to 5 X 10~ 5 g-ml ' over 24 hours. Addition of acetylcholine chloride to the same 
final concentration had no detectable effect. These findings suggest that nerves are 
not involved in the tunic responses, as peripheral nerves in tunicates typically operate 
through cholinergic synapses (e.g., Florey, 1967; Mackie et ai, 1974). 

Electrical activity ofascidiozooids 

Recordings from the zooids show ciliary arrest potentials (CAPs) like those de- 
scribed in numerous other tunicates (reviewed by Bone and Mackie, 1982). As re- 
ported by Mackie (1974) for another compound ascidian, Distaplia occidentalis, the 
CAP patterns of different zooids in the colony show no coordination. Attenuated 



198 



G. O. MACKIE AND C. L. SINGLA 




FIGURE 8. Cloacal aperture before (A) and after (B) stimulation of the tunic. Three TPs were elicited 
10 seconds apart, leading to reduction of the circumference of the aperture by 17%. 



CAPs can be recorded a short distance down the zooid stalk and in the upper sheet 
of the tunic close to the zooids; these signals are probably picked up electrotonically, 
rather than being conducted events. 

Recordings from the vascular ampullae show small potentials similar to those 
recorded from the ampullae of colonial styelids and Perophora, and like them exhib- 
ited in a rhythm coinciding with the contractions which propel blood through the 
system (Mackie and Singla, 1983). Ampullae belonging to the same zooid are coordi- 
nated, but those of different zooids are not. The two ampullae shown in Figure 2B 
belong to different zooids and are out of phase. Cycle time is about 140 s and, as in 
Botrylloides, the potentials typically occur in doublets. 

Effector correlates of tunic pulses 

So far as we know, tunic pulses have no relationship to the electrical pulse patterns 
recorded from the zooids, and vice versa; nor do TPs seem to be involved in the 
locomotory process. Locomotion has been observed in colonies showing no TP pat- 
terns as well as in those showing such patterns. In fact, it seems unlikely that locomo- 
tion is controlled by any colony-wide coordinating system. The pulsatile movements 
of the blood vessels and vascular ampullae certainly play a part in locomotion but 
they are not coordinated on a colonial basis. 

The only clearly demonstrable effect of TP activity is the contraction of the cloacal 
apertures (Fig. 8). Constriction of the aperture results in an increase in the rate of 
water flow through the opening. This occurs in a stepped manner, with each step 
corresponding to a single TP (Fig. 9). Following cessation of TPs, the aperture relaxes 
slowly. This effect of TPs can be observed both during experimentally induced and 
spontaneous TP activity, given repetitive firing at a sufficiently high frequency. 

For more detailed study, given the sluggish nature of the response, it was conve- 



TUNIC RESPONSE SYSTEM 



199 




FIGURE 9. Change in rate of water flow through a cloacal aperture following stimulation of the tunic. 
A stimulating electrode (S) on the tunic evokes TPs, picked up with a recording electrode (R) and shown 
as small events following large stimulus artifacts on upper trace. Lower trace shows stepped increase in 
flow rate accompanying the stimulus train, recorded with a glass based thermistor flow meter (F). Following 
the stimulus train, flow rate returns to normal as the cloacal aperture dilates. 



nient to monitor changes in the cloacal apertures visually, using a scalar eyepiece to 
measure diameters, from which changes in circumference could be calculated. (The 
myocytes are arranged in circular arrays around the openings, so changes in circum- 
ference represent length changes in the contractile tissue. ) As expected, long TP bursts 
produce more contraction than short TP bursts at any given pulse frequency. With 
shocks set to evoke TPs at intervals of six seconds, summation of contractions is 
approximately linear until the preparation has shortened to about two-thirds of its 
resting length, when the curve flattens out (Fig. 10). Pulses more than about 15 sec- 
onds apart do not usually produce a summing response. It was observed that relax- 
ation following contraction generally involves a period of hyperextension, after which 
the preparation returns to its resting length (Fig. 1 1 ), but no TP activity accompanies 
this final phase. Finally, it was shown that with long duration pulse trains at any 
given frequency, the preparation fails to maintain the level of contraction exhibited 
initially, but lengthens to a plateau level which is maintained indefinitely (Fig. 12). 



100 



I 

g 

5 



8 

u 



50 



5 10 

Number of pulses at 10 p.p.m. 



15 



FIGURE 10. Summation of contractions during TP trains evoked by stimulation at 10 pulses per 
minute. 



200 



G. O. MACKIE AND C. L. SINGLA 



o> 
o 

CO 



3 

E 



o 

^ 

'o 



last pulse 




a 3 pulses 

1 5 pulses 



-I 



time (mins) 



FIGURE 1 1 . Changes in circumference of a cloacal aperture following TP trains of 3 pulses and 1 5 
pulses, both at 10 pulses per minute. 



For the experiments reported above, stimulation parameters were deliberately 
kept at a moderate level, so that each shock produced a single propagated TP. 
Stronger stimulation which causes multiple firing of the TP system, or repetition of 
normal stimuli at higher frequencies can produce almost complete closure of the 
apertures. Under these circumstances, the whole upper surface of the tunic has con- 
tracted to some extent, and the cloacal space has diminished. Therefore, although no 
attempt was made to quantify these observations, it seems clear that the contraction 
of the cloacal apertures is only part of an overall contractile response involving the 
whole tunic. 

Re-examination of Botryllus 

The discovery of a tunic conduction system in Diplosoma raised questions about 
our earlier results with Perophora and with Botryllus and its relatives (Mackie and 



E 
o 

5 



control 




Time (mins) 

FIGURE 12. Changes in circumference of a cloacal aperture as observed over a five minute period 
with stimulation at two different frequencies, and with an unstimulated control. Each shock produced a 
single TP. 



TUNIC RESPONSE SYSTEM 201 

Singla, 1983), where we found that coordination of colonial activities occurred by 
epithelial conduction in the blood vessels connecting the zooids. It is conceivable that 
in these cases conduction also involves myocytes in the tunic itself. Therefore, the 
earlier investigation was repeated using B. schlosseri, which grows on the walls of the 
storage tanks at the Naples aquarium. The earlier results were correct. The propa- 
gated signals in Botryllus can be recorded only from the vascular ampullae and blood 
vessels. There is no sign of conduction in parts of the tunic where there are no blood 
vessels. It was confirmed that the blood vessel impulses ("network potentials," NPs) 
cause ciliary arrests in the zooids, as earlier claimed. Therefore, this NP system in 
Botryllus is distinct from the TP system in Diplosoma. It is interesting that Diplosoma 
has a version of the NP system, but it operates only within the confines of individual 
zooids, and presumably functions to coordinate the contractions of the four vascular 
branches which run out into the tunic from each zooid. Thus, the NP system occurs 
in Aplousobranchs (Diplosoma), Phlebobranchs (Perophora), and Stolidobranchs 
(various Botryllinae) and must be regarded as a basic ascidian action system. To date 
the TP system has been identified in only one family of aplousobranchs, the Didemni- 
dae, represented in Diplosoma, and may be peculiar to this group. 

DISCUSSION 

The evidence presented here demonstrates the ability of the tunic of a didemnid 
ascidian to conduct all-or-none propagated impulses in response to electrical stimula- 
tion and for these signals to cause contractions of the tunic. No such findings have 
been reported for other species, and it seems probable that the properties of conduc- 
tion and contraction are not widespread in the Ascidiacea, and may indeed prove to 
exist only in the family Didemnidae. The system enables the colony to control its 
exhalent water stream, a function performed in most ascidians at the individual zooid 
level, by muscles in the walls of the atrial siphons. The zooids in didemnids lack atrial 
siphons, and the only way of controlling water outflow is by regulating the size of the 
openings in the tunic (the common cloacal apertures). Therefore it seems possible 
that the properties of conduction and contraction in the didemnid tunic evolved in 
parallel with the reduction and loss of the atrial siphons of the zooids, primarily as a 
way of allowing the organism to control its exhalent water currents. 

It is not clear exactly what benefits would be associated with the ability to regulate 
water flow through the colony. Strong stimulation can produce almost complete clo- 
sure of the cloacal apertures, which might be advantageous in the presence of a preda- 
tor. Less strong stimulation causes constriction of the apertures and produces narrow, 
high velocity water plumes, which rise to a greater height above the colony; this would 
reduce the amount of water recycled through the colony and increase advection of 
fresh water from the surroundings. Contractility also allows the colony to regulate 
the volume of water in its cloacal cavity thereby enabling it to expand or contract, an 
adaptation that might be put to a variety of uses. As noted earlier, Diplosoma virens, 
which possesses photosynthetic symbionts (Prochloron) in its tunic, expands and con- 
tracts on a diurnal basis (J. S. Ryland, pers. comm.). 

We have searched in vain for evidence that the tunic conduction pathway medi- 
ates protective responses of the zooids. The majority of colonial animals have some 
means of coordinating their defensive responses, and this is true of ascidian colonies 
like Perophora and Botryllus, whose zooids are coordinated by signals transmitted 
through the colonial vascular system (Mackie and Singla, 1983). But Diplosoma ap- 
pears to be an exception. Here there is no colonial vascular network and impulses 
propagated in the tunic conduction system seem to have no effect on the zooids. 



202 G. O. MACKIE AND C. L. SINGLA 

Indeed, as Delia Valle (1908) remarked in the context of locomotory behavior, the 
tunic has its own 'individuality,' meaning that it has a life of its own, functioning 
without reference to the zooids contained in it. 

Re; sg the cellular basis for conduction and contraction in the tunic, there 
can be little doubt that the cells termed myocytes are responsible for the contractions. 
It alsc >eems likely that these cells conduct the electrical signals for their own contrac- 
he only other cells arranged in a net-like configuration the filopodial cells 
lie in a different layer of the tunic and make few contacts with the myocytes, so they 
are probably not the conducting elements. There is nothing inherently unlikely in 
the idea of a primitive contractile system which conducts its own impulses. Vertebrate 
cardiac muscle and many sorts of smooth muscle show this ability. However, we are 
hestitant to call the cells in question muscle cells because they exhibit a lower level 
of differentiation than true smooth muscle cells in tunicates, both in terms of their 
general morphology and of their ultrastructure. The term 'myocyte' seems best for 
these actin-loaded cells which lack thick myofilaments, are arranged in a loose net- 
work, conduct impulses very slowly, and show very long contraction latencies. 

Non-muscle contractility is well developed in ascidians. Tail resorption in ascid- 
ian tadpoles involves the rapid transformation of squamous epithelial cells into tall, 
flask-shaped cells during which actin microfilaments become aligned in the apical 
(Distaplia) or basal (Botryllus) cytoplasm. Discussing these findings, Cloney (1982) 
states that "the caudal epidermis clearly provides the driving force in tail resorption." 
Sperm release in Ciona involves contraction of the sperm duct epithelium, again by 
organization of actin microfilaments. The assembly of the filaments is triggered by 
light (Woollacott and Porter, 1977). Microfilaments are also involved in the contrac- 
tions of the vascular ampullae of colonial styelids like Botryllus, Botrylloides, and 
Metandrocarpa (DeSanto and Dudley, 1969; Katow and Watanabe, 1978; Mackie 
and Singla, 1983). The epithelial cells in these cases communicate via gap junctions, 
which presumably provide intercellular pathways for transmission of the impulses 
which coordinate the contractions of the ampullae. A similar mechanism may apply 
to the myocyte network in the tunic of Diplosoma, but intracellular recordings and 
demonstration of coupling between myocytes are required to prove this. The possibil- 
ity that the myocytes communicate via chemical synapses cannot be ruled out, espe- 
cially in view of the system's sensitivity to magnesium. 

Thus we believe the tunic myocyte net is a system evolved de novo in Diplosoma 
and probably in other didemnids to bring about coordinated contractions of the ex- 
halent water openings, thus bringing water flow under colonial control. Contractions 
are slow, conduction velocity is the slowest on record for any animal with the excep- 
tion of hexactinellid sponges (Mackie el ai, 1983), the system has a limited carrying 
capacity in terms of impulse frequency, and it appears to fatigue very quickly. Nerves 
and muscles probably would allow the animal to respond with much more alacrity; 
however, there are no nerves or muscles in the tunic of any ascidian, so it would 
seem that there was no evolutionary starting point for a conventional neuro-muscular 
system and a completely new type of cell the myocyte had to be evolved, albeit 
as a rather inefficient substitute. 

These findings emphasize the unusual versatility of the Tunicata in developing 
mechanisms of colonial coordination without ever using simple, direct nervous inter- 
connections. Diplosoma uses an excitable myocyte network in the tunic, Botryllus 
uses its excitable vascular epithelium, the zooids in a colony of Pyrosoma signal to 
each other by responding visually to each others' biolurninescent flashes (Mackie and 
Bone, 1978), and salps relay signals between zooids by excitable epithelial pathways 
arranged in series with nerves (Bone et al, 1980; Anderson and Bone, 1980). It is 
unlikely that these examples exhaust the list of possible mechanisms. 



TUNIC RESPONSE SYSTEM 203 

ACKNOWLEDGMENTS 

The bulk of the experimental work reported here was performed during a visit by 
G.O.M. to the Stazione Zoologica, Naples, Italy, aided by travel and operating grants 
from the Natural Sciences and Engineering Research Council of Canada (NSERC). 
We are most grateful to the Director and staff of the Stazione Zoologica, and particu- 
larly to Dr. Amedeo de Santis, for making facilities and a complete set of electronic 
recording equipment available for this study. Some follow-up work was performed 
at the Friday Harbor Laboratories of the University of Washington, Friday Harbor, 
Washington, and we thank the Director, Dr. A. O. D. Willows for providing space and 
facilities there. C.L.S. thanks NSERC for salary and funding for use of the electron 
microscope facilities of the University of Victoria. Ms. Joan Glazier of the Bamfield 
Marine Station, British Columbia, collected and shipped to us specimens of Diplo- 
soma macdonaldi. 

During the preparation of this work, we have had helpful correspondence or dis- 
cussions with Dr. Stuart Arkett, Dr. Michael Cavey, Dr. Richard Cloney, Ms. Sarah 
Cohen, Dr. Jean Godeaux, Dr. John Ryland, and Dr. Thomas Schroeder. 

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Reference: Biol. Bull. 173: 205-221. (August, 1987) 



BIVALVE HEMOCYANIN: STRUCTURAL, FUNCTIONAL, AND 
PHYLOGENETIC RELATIONSHIPS 

C. P. MANGUM 1 , K. I. MILLER 2 , J. L. SCOTT', K. E. VAN HOLDE 2 , 

AND M. P. MORSE 3 

Department of Biology. College of William and Mary, Williamsburg, Virginia 23185; 2 Department of 

Biochemistry and Biophysics, Oregon State University, Con'allis, Oregon 97331; and* Marine Science 

Center and Biology Department, Northeastern University. Nahant. Massachusetts 01980 

ABSTRACT 

The hemocyanin-like molecule found in the blood of the most primitive bivalves 
(protobranchs) reversibly binds O 2 . Its respiratory properties and its sedimentation 
behavior are both distinctive. Although its electron-dense image looks like that of the 
gastropod hemocyanins, its molecular weight differs from those of all other molluscan 
Hcs and is more consistent with the concept of bivalve hemocyanin as a pair of octo- 
pod hemocyanins. Bivalve hemocyanin occurs in the solemyoids as well as the nucu- 
loids, which argues for the integrity of the Protobranchia as a natural taxon. The 
ancestral bivalve O 2 carrier was previously believed to be a simple intracellular hemo- 
globin, which is found in the less primitive Pteriomorpha. The most obvious interpre- 
tation of the present results, however, is that hemocyanin is the primitive bivalve O 2 
carrier and that it was replaced by the red blood cell, which originated at least twice: 
once in the pteriomorph bivalves and at least once in other taxa. 

INTRODUCTION 

Recently Morse el al. (1986) reported the presence of electron dense images that 
resemble molluscan hemocyanins (Hcs) in the blood of two nuculoid bivalves. In 
addition, the denatured subunits exhibited the same anomalous behavior during so- 
dium dodecyl sulfate polyacrylamide electrophoresis as those of some other mollus- 
can Hcs, viz., they migrated to a position corresponding to a lower molecular weight 
than expected from other aspects of quaternary structure (Van Holde, 1983; Ryan et 
al., 1985). Finally, copper electrons were identified in the blood of one species by X- 
ray spectroscopy of sections of the auricle. Thus the evidence indicates the existence 
of a molecule that closely resembles molluscan He in the most primitive members of 
the class Bivalvia, which was formerly believed to use either heme proteins or no O 2 
carrier at all (Mangum, 1 980a; Terwilliger and Terwilliger, 1985). 

This finding has considerable phylogenetic importance. First, the Hcs found in 
the various molluscan classes are believed to exhibit differences, albeit quite subtle 
ones, in quarternary structure (Ryan et al, 1985). Therefore a comparison may eluci- 
date evolutionary relationships between them. Second; it has been suggested that red 
blood cells (RBCs), which occur in the somewhat less primitive pteriomorphs, may 
represent the ancestral condition among the bivalves (e.g. , Mangum, 1 980a). Third, if 
instead He is the primitive O 2 carrier in the class, then the RBC must have originated 
independently on at least two occasions, within the bivalves and in other taxa. 

In the present contribution we report evidence of reversible O 2 binding, which 
demonstrates that the molecule in the blood of both groups (solemyoid as well as 

Received 2 March 1987; accepted 1 May 1987. 

205 



206 C. P. MANGUM ET AL. 

nuculoid) of protobranch bivalves is a typical O 2 carrier, not a He-like precursor. We 
also describe additional aspects of molecular structure and respiratory function that 
enable comparison of the protobranch blood O 2 carrier with the Hcs found in other 
molluscs. Finally, we explore the implications of our findings for RBC phylogeny. 

MATERIALS AND METHODS 

Acila castrensis (Hinds) and Cyclocardia (=Venericardia) ventricosa (Gould) 
were collected near San Juan Island, Washington. Yoldia limatida (Say), Nucula 
proximo. Say, and Solemya velum Say were purchased from commercial sources in 
Massachusetts. 

O 2 uptake (VO 2 ) of intact animals was determined as the depletion of O 2 in the 
PO 2 range 120-159 mm Hg, measured with a self-stirring polarographic electrode 
(Yellow Springs Instrument Co. Model 54). To prevent extraneous O 2 uptake by shell 
epibiota, the animals were disinfected by immersion for a few seconds in benzalkoni- 
um chloride (0.13%). Vacant shells given this treatment do not take up appreciable 
volumes of O 2 . 

Blood was obtained by first inducing the animals to empty their mantle cavities 
and then prying open the valves about 1 mm or less with a jeweler's screwdriver. The 
valves were reflected backwards about 270 and the animals placed in a small funnel 
draining into an Eppendorf tube. When the blood ceased to drain, additional volumes 
were obtained by centrifuging the animals at a very low speed. After repeated prob- 
lems with apparent proteolysis during sedimentation analysis, we collected the blood 
of Y. limatida by draining it directly into a mixture of protease inhibitors, which 
appeared to enhance the stability of the preparation. About 1 ml blood was drained 
into a 60 n\ solution containing 30 pg leupeptin, 30 pg pepstatin A, and 3 yumoles 
phenylmethylsulfonyl fluoride. 

The gills of Y. limatida were dissected and extracted with 0.5% Na 2 CO 3 . The 
extract was diluted by 10% with pyridine, reduced with a few grains of sodium dithio- 
nite, and, due to its very small volume, examined with Zeiss micro- and Hartridge 
reversion spectroscopes (Mangum and Dales, 1965). 

The bloods were centrifuged and immediately prepared for electron microscopy 
(Zeiss EM- 109) by negative staining with uranyl acetate (Mangum etai, 1985; Morse 
et al., 1986). In the present investigation the blood was diluted with 0.05 M Tris 
maleate buffer + 10 mMCaC! 2 (pH 7.63) by factors ranging from 1:9 to only 1:50, 
because its appearance suggested a low He concentration. The sample and the stain 
were applied to the grid with an atomizer. 

O 2 binding was determined on fresh (never frozen) blood samples from A. cas- 
trensis, S. velum, and Y. limatida by the cell respiration method (Mangum and Lyk- 
keboe, 1979). Due to the size of the N. proxima individuals (2-5 mm), it was neces- 
sary to stockpile frozen material until the requisite volume (300 n\) accumulated. 
About half of the material on which the measurements were performed had been 
frozen. The samples were diluted by 10% with Tris maleate (final concentration 0.05 
M) buffered seawater (32%o) containing commercial yeast cells. An attempt to first 
concentrate A. castrensis blood by membrane filtration was unsuccessful due to pre- 
cipitation of some of the material, which was also noted during the O 2 binding mea- 
surements. No precipitation of the Y. limatula, S. velum, or N. proxima material was 
observed. 

The O 2 affinities of heme proteins are often concentration dependent. Because 
most experimental procedures require dilute solutions, the results do not accurately 
reflect in vivo values. To obtain physiologically meaningful data for the branchial 



BIVALVE HEMOCYANIN 207 

heme protein of Y. limatula, O 2 binding in the present investigation was also deter- 
mined using whole gills, dissected intact. An absorption spectrum of the bathing me- 
dium indicated that there was no loss from the gills during the measurement. It was 
necessary to modify the cell respiration procedure because the method requires that 
the rate of free O 2 depletion be linear. This is achieved by lowering the PO 2 with 
particles such as isolated mitochondria or yeast cells which are so small that the diffu- 
sion distance is not limiting. If the O 2 uptake of whole unperfused gills had been 
allowed to make an appreciable contribution to total O 2 uptake, this condition would 
clearly have been violated and the apparent O 2 affinity would have been erroneously 
low due to an extraneous departure from linearity. The problem was circumvented 
by first determining the VO 2 of the gills and then adding large numbers of yeast cells 
so that yeast VO 2 was more than 10 times gill VO 2 . The cell respiration method also 
requires that the rate of O 2 depletion be slow enough to permit equilibration of the 
electrode at any PO 2 . If this condition had been violated, the result would have been 
an erroneously high O 2 affinity, because an apparent oxygenation state would have 
coincided with a PO 2 that actually had obtained earlier, at a higher oxygenation state. 
The possibility was eliminated by ascertaining equilibration under the following ex- 
perimental conditions: using the He of the crab Cancer magister, whose respiratory 
properties are well known (e.g., Graham el ai, 1983), the rate of O 2 depletion was 
increased until an erroneously high O 2 affinity was obtained. The period for depletion 
of free O 2 from 100 to 0% air saturation was considerably less than 25 s. In the mea- 
surements on gills, much longer periods (87-233 s) were employed. 

Absorption spectra of the medium and of fresh Hcs were determined with Beck- 
man DK-2A and Varian 2200 spectrophotometers. To eliminate light scatter, the He 
samples were first diluted with dissociating buffer (0.05 MTris HC1, pH 8.95 + 0.05 
M EDTA) by 50 to 97% depending on color intensity. 

All sedimentation experiments were performed in a Beckman Model E analytical 
ultracentrifuge equipped with scanner optics. Wavelengths in the vicinity of the He 
bands at 280 and 345 nm were used. Temperature was controlled to <0.1C. Sedi- 
mentation coefficients were measured from the midpoints of the well-defined bound- 
aries and corrected to S 20 . w in the usual way. The sedimentation equilibrium experi- 
ment was conducted at 1 500 rpm, using the heavy J rotor. Equilibrium was attained 
when no difference could be noted between scans approximately 6 h apart. After 
equilibrium the rotor was accelerated to 6000 rpm and a baseline recorded approxi- 
mately 4 h later. 

RESULTS 
O 2 uptake 

Intact individuals of Acila castrensis take up O 2 measurably, but VO 2 is more 
than two orders of magnitude lower than in the heterodont Cyclocardia ventricosa 
(Table I), which was collected from the same bottom on the same occasion and held 
in the laboratory in the same container for the same period. The difference in body 
size can account for only a small fraction of the difference in O 2 uptake. Moreover, 
VO 2 is also orders of magnitude lower in A. castrensis than in the pterimorph Noetia 
ponderosa, a much larger animal measured at a slightly lower temperature (Table I). 

Absorption 

The dissociated subunits of the four bivalve Hcs absorb at 280 and 345 nm (Fig. 
2). Other molluscan Hcs absorb in the same regions (Nickerson and Van Holde, 



208 



C. P. MANGUM ET AL. 



TABLE I 

Oxygen uptake in bivalves with specialized oxygen carriers 



Species 


O 2 carrier 


VO 2 

(Ail/g dry wt-h) 


Dry wt. 

(mg) 


Temp. 
(C) 


Source 



Ad la cti 
Vene 

' "iricosa 



He 
extracellular Hb 



Noeliaponderosa intracellular Hb 
Glycimeris 
nummaria intracellular Hb 



2.0 0.9(6) 107.9-122.8 11.5 present data 

269.5 15.5(6) 33.5-34.9 
148 5 X 10 3 

ca. 49. 5 a 2.7X10 3 +16-20 Kruger, 1957 



11.5 present data 

Deaton and Mangum, 
10 1976 



1 Converted from original data assuming that dry wt. = 20% wet wt. Mean SE (n). 



1971). R. C. Terwilliger kindly communicated data for polyplacophoran He, which 
have not been reported in the literature. Observations on the He of the chiton Chae- 
topleura apliculata were also made together with those reported here. The peak at 
345 nm disappears in the presence of sodium borohydride (e.g., Fig. 1). 

Extracts of the gill of Y. limatula clearly form a pyridine hemochromagen with 
absorption bands at 542 and 556 nm. High concentrations of red granules also were 
observed in the nerve ganglia and connectives of this species but not in A. castrensis or 
N. proxima, which appeared to lack branchioglobin (Bb) as well. However, a seawater 
extract of the whole bodies of N. proxima appeared to form a pyridine hemochroma- 




320 



340 



3 50 



FIGURE 1. Absorption spectra of protobranch Hcs. (A) Acila castrensis, (N) Nucula proxima, and 
(S) Solemya velum. Abscissa is wavelength in nm and ordinate is absorbance. 



BIVALVE HEMOCYANIN 



209 



B 



100 



10 
80 

60 
4.0 



e 2.0 

E 



1.0 
0.8 

0.6 




10 




\ 



0.1 



70 



72 



7.4 76 

P H 



7.8 



8.0 



0.01 



o 

o 



s 



70 



7.2 



7.4 



7.6 
PH 



78 



80 



82 



1 


1 


1 


0.1 


1 


10 




p o 2 





FIGURE 2. A. PH dependence of O 2 binding by Acila castrensis (), Nucula proximo. (D), Solemya 
velum (<>), and Yoldia limatula (O) Hcs. 20C, 0.05 M Tris maleate buffered blood. B. Hill plot of 6 2 
equilibrium of Yoldia limulata (, pH 8. 10.) and Acila castrensis (O, pH 7.78) Hcs. 



gen, although the visual observation could not be confirmed even by microspectro- 
scopic observation due to the very small volume (ca., 10 n\) obtained. 

O 2 binding of the bloods 

Unlike other Hcs, A. castrensis He binds O 2 non-cooperatively (e.g., Fig. 2). The 
Hill coefficient ( 50 ), which is independent of pH, is 1 .03 (0.03 SE, n == 11). Among 
the Hcs, A. castrensis also has an unprecedentedly high O 2 affinity (Fig. 2), thus re- 
sembling tissue O 2 carriers more than most blood O 2 carriers. Unlike tissue O 2 carri- 
ers, however, A. castrensis He has a small but significant normal Bohr shift. The slope 
of the regression line describing the data in Figure 2, or A log P 5 oM pH, is -0.23 
(0.08 95% C.I.)- HcO 2 binding in A", proxima (which belongs to the same family 
as A. castrensis) is also non-cooperative (0.94 0.09 SE; n = : 7) and it has a similar 
Bohr shift (-0. 16 0.07 95% C.I.) although its O 2 affinity is somewhat lower. The 
He of Y. limatula (which belongs to a different family) has a much lower O 2 affinity, 
though still fairly high for a molluscan He, and it is moderately cooperative (Fig. 2). 
Its Bohr shift is indistinguishable from that of the other nuculoid Hcs (-0.24 0.05). 



210 C. P. MANGUM ET AL. 




FIGURE 3. Electron micrographs of bivalve Hcs. A. Yoldia limatula. B. Solemya velum. Scale bar 
50 nm. 



The He of S. velum (which belongs to a different order) resembles Y. limatula He in 
terms of O 2 affinity and cooperativity, but its Bohr shift is much larger (-0.6 1 0. 1 7). 
O 2 carrying capacity of the bloods (HcO 2 + O 2 ) was estimated from absorbance 
at the active site, using the extinction coefficient for Busycon He (Nickerson and Van 
Holde, 1971). At 1 1.5C and 32%o salinity the value for A. castrensis blood is 1.05 
ml/ 100 ml, for S. velum is 1.00 ml/ 100 ml, and for one sample from Y. limatula is 
0.96 ml/ 100 ml. N. proxima blood, which is much bluer than the others, carries 2.85 
ml/ 1 00 ml. The figure for A. castrensis should be regarded as low due to precipitation 
in the sample. However, a similarly low figure (0.76 ml/ 100 ml) for another sample 
from Y. limatula was obtained from integrals of the curves describing deoxygenation 
(procedure detailed by Mangum and Burnett, 1986). Moreover, the difference be- 
tween N. proxima, S. velum, and Y. limatula cannot be due to starvation of the latter 
two in the laboratory (which, in fact, has not been reported for molluscan Hcs) since 
they were held for the same period (<2 days). 

O 2 binding by gills 

Two determinations of O 2 binding by intact Y. limatula gills, which should pro- 
vide physiologically meaningful information, gave P 50 values of 0.43 and 0.46 mm 
Hg and n values of 0.98 and 1 .02 (20.3C, ambient pH 8.0 1 ). 

Electron microscopy 

Since N. proxima is so closely related to A. castrensis, the small amount of mate- 
rial available was used for other purposes. The shapes of S. velum and Y. limatula 
Hcs (Fig. 3) are indistinguishable from that of A. castrensis He, which was described 
earlier (Morse el al, 1 986). All three molecules are six-tiered cylinders and, like many 
gastropod Hcs, appear as circles in top view and as squares in side view (see Ghiretti- 
Magaldi et al., 1979, van Bruggen el al, 1981). They lack the "belt," or unequal 
spacing of the six tiers, found in one species (van der Laan el al., 1981). The width 
(31 nm) of the Y. limatula squares appears to be slightly but significantly (P < .001 
according to Student's / test) smaller than that of 12 tiered cylinders found in the 
blood of the gastropod Busycon canlicutatum. These dimensions were determined by 
mixing a small volume of B. canaliculatum blood with a large volume of Y. limatula 



BIVALVE HEMOCYANIN 



211 




CcTMg* 




EDTA 




YOLDIA 



T T T 
15S 96S 

AC I LA 



FIGURE 4. Scanner traces showing dissociation and reassociation of bivalve Hcs at pH 7.65, 20C. 
Yoldia limanda: ( 1 ) in 0.05 M Tris-HCl, 50 mM MgCL, 10 mM CaCl 2 ; (2) dialyzed against 0.05 A/Tris- 
HC1 + 10 mM EDTA; (3) dialyzed back again against the original buffer. Acila castrensis: (4) as in 1; (5) 
as in 2; (6) as in 3. In 5 and 6 the middle boundary sediments at about 55S. 



blood and then measuring the width of all (24) of the 12-tiered cylinders observed 
and a sample of 100 6-tiered cylinders. The bivalve circles have a five-fold rotational 
symmetry and a collar and a cap. When dissociated to halves, the molecule looks like 
a three-tiered rectangle, which absorbs more stain at one end than the other, and as 
circles, only some of which have collars and caps (see Fig. 3 in Morse et ai, 1986). The 
images of half molecules, which have also been described for Helix He, are believed to 
reflect the absence of collars and caps at the broken surfaces (van Bruggen et ai, 
198 1 ). Like gastropod Hcs, the bivalve squares are about 35 nm long. 

Physical characterization 

Sedimentation velocity experiments with A. castrensis and Y. limatula Hcs were 
performed at room temperature and under a variety of solvent conditions. The results 
are summarized as follows: 

(1) In 0.05 M Tris-HCl buffer (pH 7.65) containing 50 mMCaCl 2 and 10 mM 
MgCl 2 , both Hcs exhibited single, sharp boundaries. The sedimentation coefficients 
(S2o,w), when corrected to standard conditions and extrapolated to zero He concen- 
tration, were 95.8 for A. castrensis He and 88.8 for Y. limatula He. 

(2) When the two He solutions were dialyzed exhaustively against 10 mMEDTA 
in 0.05 M Tris-HCl (pH 7.65), they behaved differently (Fig. 4). Y. limatula He disso- 
ciated completely to yield a single boundary with S 20 .w : 15.9S. Under the same 
conditions A. castrensis He showed incomplete dissociation, yielding two compo- 
nents with S 20 ,w ~-~- 54 and 18S. Attempts at reassociation also gave quite different 
results. Upon dialysis of the Tris EDTA treated material back to Tris Ca +:: + Mg +2 , 
Y. limatula He quantitatively reassociated to the 89S component. Under the same 
conditions only partial reassociation could be attained with A. castrensis He. 

(3) When Y. limatula He was dialyzed against a series of dilutions of the Tris 
buffer in which the divalent cations were reduced to '/ 10 , '/ 20 , '/to, and finally Vioo of 



212 



C. P. MANGUM ET AL. 



1001 



50- 




5.0 mM MgCl2 
1.0 mM CaCI 2 

FIGURE 5. Relative amounts of Yoldia limatula He of three aggregation states when equilibrated to 
0.05 A/Tris buffers (pH 7.65; 20C) containing varying amounts of MgCl : and CaCl 2 (see text for details). 



their original concentrations, the resultant dissociation yielded a mixture of three 
components: the 89S He (the whole molecule), another with a sedimentation coeffi- 
cient of 55S (probably a half molecule), and a third with a sedimentation coefficient 
of about 15S (Fig. 5). The present data do not indicate just what multiple of the 
polypeptide chain this smallest product represents, but the sedimentation coefficient 
corresponds to that of the dissociation product in the presence of EDTA. 

The reversible dissociation behavior of Y. limatula He strongly suggests that, like 
Octopus He (Van Holde and Miller, 1985), it is composed of a single type of subunit. 
In contrast, the incomplete reassociation of A. castrensis He is more like that of other 
molluscan Hcs(Van Holde and Miller, 1982). Furthermore Y. limatula He, like Octo- 
pus He, dissociates in the presence of EDTA at a much lower pH than normally 
required for other molluscan Hcs. Although divalent cation levels must be reduced 
to extremely low levels before dissociation begins, Y. limatula He dissociates at pH 
7.65, which is probably close to the physiological value. In all likelihood, at higher 
pH it would dissociate at higher divalent cation levels. 

DISCUSSION 
Respiratory properties and their relationships to protobranch biology 

Allen (1978) suggested that protobranch bivalves are able to exist with their small 
and, in his view, relatively inefficient feeding organs because they have low metabolic 
rates. The present findings support his suggestion, at least with respect to aerobic 
metabolism. However, we should point out that, relative to bivalves that both use the 
gill as a filter-feeding organ and lack an O 2 carrier, the branchial surface area is also 
small in thepteriomorph Noetia ponderosa, whose feeding has not been investigated 
and whose VO 2 is not especially low (Mangum, 1980a). Almost certainly VO 2 is in- 
fluenced by other factors in addition to feeding efficiency. While it is believed that 
conventional feeding in Solemya is supplemented or perhaps even supplanted by 
a symbiotic relationship with chemoautotrophic bacteria (Cavanaugh, 1980, 1983; 
Felbeck, 1983; Doeller, 1984; Fisher and Childress, 1984; Reid and Brand, 1986), no 
sign of bacteria can be found in electron micrographs of the gills of protobranchs such 
as A. castrensis (mentioned by Reid and Brand, 1986) and Y. limatula (M. P. Morse, 
unpub. obs.). 



BIVALVE HEMOCYANIN 213 

The uniformly normal Bohr shift of the bivalve Hcs resembles those of polyplaco- 
phoran and cephalopod Hcs. Gastropod Hcs have either reversed Bohr shifts (proso- 
branch), a combination of reversed and normal Bohr shifts (prosobranch and pulmo- 
nate), or none at all (opisthobranch). As indicated above, the extremely high O 2 
affinity and lack of cooperativity of A. castrensis He is unique. The moderate cooper- 
ativity and O 2 affinity of S. velum and Y. limatida Hcs are common among the mol- 
luscs, although examples of much greater cooperativity are known (Mangum, 
1980b). O 2 carrying capacity appears distinctively low, at least in S. velum and Y. 
limatida, but typical of molluscan HcO 2 transport systems in N. proxima. Why the 
nuculoids, with such similar respiratory and cardiovascular systems, should have Hcs 
with such different respiratory properties remains to be elucidated. 

The anatomical relationships between the protobranch O 2 carriers also are in- 
triguing. At least in Y. limatida and S. velum, O 2 must move from the environmental 
source into the heme protein-containing branchial epithelium. From there the O 2 
moves into the He-containing blood, where it is carried by convection to the meta- 
bolic sink. But in both species the O 2 affinity of the branchioglobin (Bb) is higher than 
that of the He (see Doeller et ai, 1983, for values for S. velum Bb). The arrangement 
violates the fundamental design principle of an O 2 transfer system, which mandates 
the highest O 2 affinity in the compartment most remote from the environmental 
source. Bb must actually be a barrier to O 2 influx as long as it is not fully oxygenated. 

The physiological question is complicated by uncertainty surrounding the func- 
tion of bivalve Bb and other tissue heme proteins. Doeller (1984) suggested that S. 
velum Bb transports sulfide to the chemoautotrophic bacteria in the gills; the sulfide- 
oxidizing bacteria are believed to serve as key components of a newly discovered 
mode of animal nutrition. As pointed out by Dando et al. (1985), this function does 
not preclude the possibility of others, such as facilitated diffusion or O 2 storage. 

We noted that the period from onset to completion of nonlinearity of O 2 uptake 
by Y. limatida gills (in the absence of yeast cells) was only 162 s. This period includes 
both the diffusion-limited and BbO 2 -supplied components. The molecule cannot be 
an O 2 store of significant longevity. We suggest that BbO 2 carrying capacity also be 
considered in the continuing debate on functions of tissue heme proteins. 

Structural properties 

The sedimentation coefficients observed for the bivalve Hcs are surprisingly low. 
The Hcs that would seem to resemble bivalve He in shape are the six-tiered cylinders 
found in the prosobranch whelk Kelletia and the pulmonate snail Helix (van Bruggen 
et al., 1981). Opisthobranch and the other prosobranch Hcs studied tend to form 
larger aggregates or have special features such as unequal spacing of the six tiers. 
Cephalopod and polyplacophoran Hcs are three-tiered cylinders. 

As Table II shows, however, almost all reliable measurements of the sedimenta- 
tion coefficients of the gastropod six-tiered multiple yield values of S 20 , w between 100 
and 105S. We were struck by the value for Y. limatida He, which is 10-15% lower. 
A lower value might be explained by either a looser quaternary structure, greater 
hydration, or a lower molecular weight. The molecular weight obtained from Figure 
6 depends on the value assumed for the partial specific volume (i>) (see Van Holde, 
1985). Unfortunately we have neither an experimentally determined value for i for 
Y. limatida He nor an amino acid composition, from which it might be estimated. 
Values reported for molluscan Hcs range from about 0.73 (gastropod) to 0.74 (cepha- 
lopod). The former yields a molecular weight of 6.5 X 10 6 and the latter yields 6.8 
X 10 6 . Either is much lower than the values reported for the gastropod 100-105 
S Hcs (Table II). 



214 



C. P. MANGUM ET AL. 



TABLE II 

Comparative properties of native hemocyanin molecules oj bivalves and gastropods 



Species 


S 
20,w 

(svedbergs) 


M 
(g/molx 10 6 ) 


V 

(cm-Vg) 


Class: Gastropoda 1 








A rchachalina marginata 


102.3 


9.1 





Buccinitni undatum 


101.1 


9.0 





Bus\ -con canal iculat um 


103.2 


8.8 


.727 


Helix pomatia (at) 


104.3 


8.7 


.727 


Helix pomatia (ft) 


105.8 


9.0 





Murex trunculus 


102.7 


8.9 





Paludina vivipara 


102.5 


8.7 





Pila leopoldvillensis 


101.2 


8.7 





Class: Bivalvia 2 








Acila castrensis 


95.8 








Yoldia limatula 


88.8 


6.5-6.8 






' Data from Van Holde and Miller ( 1 982). Original references are given therein. 
2 Present data. The value of M for Yoldia hemocyanin depends upon whether a value of 0.73 (as for 
gastropods) or 0.74 (as for cephalopods) is assumed. 



The data are more consistent with the concept of bivalve He as a pair of cephalo- 
pod Hcs. A pair of polyplacophoran Hcs would have a much higher molecular weight 
(Ryan et ai, 1985; Herskovits et ai, 1986). The cephalopod 51-60S particles, how- 
ever, do not pair to form six-tiered cylinders. If they did, they might give rise to a 
particle with a sedimentation coefficient of about 90S and an electron-dense image 
much like that of Y. limatula He. If we assume that native Y. limatula He is a 20- 
mer of polypeptide chains, like other 6-tiered molluscan Hcs, then the chain weight 
must be approximately 3.4 X 10 5 . This is very close to the weight of octopod chains 



In A 



49 



5O 



51 



FIGURE 6. Determination of molecular weight of Yoldia limatula He by sedimentation equilibrium. 
A represents concentration in arbitrary units of A 345 nm; r is the distance from center of rotation. 



BIVALVE HEMOCYANIN 215 

(Gielens et al, 1986; Lamy et ai, 1986), and substantially smaller than that of gastro- 
pod chains (4.0-4.5 X 10 5 ). Such a conclusion seemingly contradicts the observation 
that bivalve chains (Morse et ai, 1 986) run more slowly than cephalopod chains (Van 
Holde and Miller, 1 982) on SDS gels. However, SDS gel electrophoresis is notoriously 
unreliable for glycoproteins such as Hcs. It has been frequently reported that esti- 
mates of subunit molecular weight are in substantial error for these proteins. 

On the basis of the available information we suggest that the bivalve Hcs may 
resemble octopod Hcs in having a small (relative to other molluscan Hcs) subunit, 
but share with gastropod Hcs the capacity to associate to 20-mers. This conclusion 
is supported by the apparently smaller width of the bivalve cylinders. In possible 
contradiction, however, we should mention that Ellerton and Lankovsky (1983) re- 
ported a 26-30 nm wide and 28-34 nm long He in the primitive archaeogastropod 
Haliotis iris. 

Phylogeny of the molluscan Hcs 

The most recent discussions of molluscan phylogeny suggest two major phyletic 
lines leading from the postulated ancestor, which is in turn descended from an acoelo- 
mate animal at the turbellarian-nemertine level of organization (Runnegar and Po- 
jeta, 1986; Salvini-Plawen, 1986). One of these phyletic lines is an aplacophoran- 
polyplacophoran lineage and the other leads through the monoplacophorans to the 
gastropods, the rostroconch-bivalve-scaphopods, and the cephalopods (Fig. 7). In 
view of the equally recent conclusion that the nemertines are, in fact, descendants of 
an annelid-like coelomate (and, we suggest, RBC-containing) animal (Turbeville and 
Ruppert, 1 986), the condition of the coelom in the ancestral mollusc probably should 
be reconsidered. Regardless, the present findings emphasize the importance of ascer- 
taining the properties of O 2 carriers (if any) in the poorly known molluscan classes 
such as the monoplacophorans and the scaphopods. We also look forward to the 
results of the sedimentation equilibrium studies of archaeogastropod He which were 
underway at the time of Ellerton and Lankovsky's 1983 report. The elucidation of 
the structures of the Hcs (if any) in these three groups and also additional members 
of other molluscan groups may have important implications for molluscan phylog- 
eny. In our view, the detail available at present does not permit a very confident 
conclusion concerning the evolutionary relationships of the molluscan Hcs. 

Origins of the red blood cell and its simple hemoglobins 

If the status of postulated transitional group is disregarded, there is some consen- 
sus among molluscan systematists concerning the ancestry of the bivalves. Along with 
the Gastropoda, Scaphopoda, and Cephalopoda, the class Bivalvia is believed to be 
descended from the Monoplacophora (Cox et al., 1969; Newell, 1969; Stasek, 1972; 
Pojeta, 1978; Runnegar, 1978; Runnegar and Pojeta, 1986; Salvini-Plawen, 1986). 
There is considerable disagreement, however, on the relationships of the different 
groups of bivalves. Newell (1969) described six subclasses and assigned the solemy- 
oids and the nuculoids to separate ones. However, Allen (1986) argued for the integ- 
rity of the Protobranchia as a subclass that includes all bivalves with simple, pectinate 
gills and described only one other subclass, the Lamellibranchia. Our findings 
strongly support Allen's (1986) view. While we cannot provide evidence for the ab- 
sence of He in all lamellibranchs, we can provide evidence of the absence of any O 2 
carrier in the blood of one pteriomorph (Modiolus demissus) and two heterodonts 
(Crassostrea virginica and Rangia cuneata): When measured with a Lexington In- 



216 



C. P. MANGUM ET AL. 



chordates 
(o) 



arthropods 



echinoderms 
(o) 




hemichordates 



Dhoronids brachiopods 





nematodes 

rotifers 
common / / 

ancestor// gastrotrichs 
platyhelminths 



ctenophores 
cnidarians 




ancestral 
eumetazoan 



B 



opisthobranchs pulmonates coleoids nautiloids 

7" 



prosobranchs 



ammonoids 

T 



higher lamellibranchs 

O 

pteriomorphs 

I o 

protobranchs 



primitive gastropods primitive cephalopods primitive bivalves 




scaphopods 
? 



polyplacophorans. 



monoplacophorans 
? 



aplacophorans 

,o 



ancestral mollusc 

?o 

FIGURE 1. Phylogeny of: A. The red blood cell; B. The molluscan O : carriers. () symbolizes RBCs 
and () symbolizes molluscan He. Question mark indicates uncertainty. 



BIVALVE HEMOCYANIN 217 

struments Co. analyzer, the total O 2 contents of these bloods did not differ from that 
of the seawater to which the animals were acclimated. Moreover, these bloods also 
lacked absorption maxima in the region of 345 nm, as did plasma of the RBC-con- 
taining pteriomorph Noetia ponderosa (C. P. Mangum, unpub. obs). 

The higher bivalve taxa, including the Heterodonta (which contains most of the 
familiar species), are regarded as suspect (Newell, 1969). But the Pteriomorpha, a 
relatively primitive group consisting of the anisomyarians, the extinct cyrtodonts, 
and the RBC-containing arcoids, appears to be a natural taxon. In addition, there is 
general agreement that the protobranch bivalves are even more primitive than the 
pteriomorphs (Newell, 1969; Allen, 1978, 1986). This relationship has several im- 
plications for the question of the origin and further evolution of O 2 transport systems, 
because it is among the pteriomorphs that one finds RBCs resembling counterparts 
and containing Hbs similar to those in other phyla at comparable levels of organiza- 
tion. The subject is of such importance that it is discussed in detail below. 

Nucleated RBCs containing either monomeric or oligomeric Hbs are found in 
seven animal phyla (summarized by Mangum and Mauro, 1985), including five 
(Phoronida, Annelida, Echiura, Nemertina, and Mollusca) that are often regarded as 
not too distantly related to one another and at an intermediate stage of phylogenetic 
development (Fig. 7A). While the limited anatomical information indicates the possi- 
bility of some distinctly different features of the RBCs in each group, it also indicates 
many similarities. The physiological information, also limited, indicates a similar 
metabolic organization of at least annelid and molluscan RBCs, which differs from 
that of avian and mammalian RBCs and, possibly, the sipunculid pink blood cell 
(Mauro and Isaacks, 1984; Mangum and Mauro, 1985). 

Within the molluscs, RBCs occur widely in one order of pteriomorph bivalves (as 
well as in a single species of heterodonts; Terwilliger et al., 1983), and they almost 
certainly occur in the Aplacophora, which was regarded by Hyman (1967; p. 69) as 
the "genuinely primitive" molluscan class (Fig. 7B). Hyman (1967; p. 65) noted that 
"the coelomic fluid has a reddish hue invested in the corpuscles, except in the Chae- 
todermidae, where the red substance, not proved to be hemoglobin, is dissolved in 
the fluid itself." Despite the caveat, it is highly likely that this pattern reflects yet 
another instance of O 2 transport by RBCs (see also Baba, 1940) as the primitive con- 
dition and of multidomain, extracellular Hb as the derived condition. Well known 
examples include the annelids as well as the lamellibranch bivalves. RBCs contain- 
ing simple heme proteins are found in more primitive species and extracellular Hbs 
that differ fundamentally from one another as well as from the simple heme proteins 
occur in more advanced taxa. One pteriomorph bivalve, believed to represent the 
transitional stage, has both kinds of Hbs in its RBCs (Grinich and Terwilli- 
ger, 1980). 

There is a strong possibility that the nemertines exhibit the same trend. Hyman 
(1951; p. 490) believed that the red color of the blood "resides in the corpuscles," 
which is true of a few marine species (Vernet, 1979). In support of this contention 
Hyman ( 195 1 ) cited the 1 872 report by Lankester, whose words indicate otherwise: 
"the colour is due to Haemoglobin diffused in the liquid" (p. 73, italics ours). Polu- 
howich (1970; also pers. comm.), who reported Hb in freshwater (and therefore not 
primitive) nemertines, did not detect RBCs. Outside of the vertebrates, RBCs are 
unknown in freshwater animals, which is believed to be due to their osmotic fragility 
(Mangum, 1980a). 

On the basis of the distribution of the RBC summarized above and illustrated in 
Figure 7, there is no compelling reason to postulate more than one origin of the 
RBC and its simple hemoglobins. One need only to suppose that the RBC originated 



218 C. P. MANGUM ET AL. 

shortly after circulating body fluids arose (Fig. 7). In more advanced groups it was 
repeatedly replaced by extracellular O 2 carriers due to the greater viscosity of the large 
bore tubes that dominate primitive cardiovascular systems (Mangum, 1976), and it 
was inherited by two more advanced deuterostome groups: the echinoderms and the 
chordates. 

The most obvious interpretation of the existence of He in the blood of proto- 
branch bivalves is that the hypothesis of a common origin of the RBC is incorrect. 
This interpretation has the following implications: Protobranchs represent the ances- 
tral bivalve condition and their HcO 2 transport system was either lost (anisomy- 
arians) or replaced (arcoids) in the pteriomorphs by an intracellular HbO 2 transport 
system of de novo origin. This interpretation is consistent with the presence of neuro- 
and branchioglobin in the same individuals that contain He. It is also consistent with 
the recent finding that the tertiary structures of the simple Hbs found in the annelids, 
bivalves, and primitive vertebrates are similar to one another and to mammalian 
myoglobin, and different from that of higher vertebrates Hbs (Perutz, 1985; Royer et 
ai, 1985; W. E. Love, pers. comm.). According to this multiple origin hypothesis, 
structural similarity of the simple Hbs of the lower animals is due to two separate 
origins from their tissue heme proteins, which also have the same tertiary structure 
(presently unknown, but probable), not a common origin. 

The weaknesses of this interpretation include uncertainty about the integrity of 
the class Bivalvia(McAlester, 1966;Cox^/. ai, 1969; Newell, 1969; Runnegar, 1978) 
and the absence of a clear selection pressure for the replacement of He with RBCs. 
The simple heme proteins found in pteriomorph RBCs would seem to offer no clear 
advantages over protobranch Hcs. Indeed, their respiratory properties are not nearly 
as plastic, at least under physiological conditions (Mangum, 1980a), and therefore 
the selection pressure would seem to be negative. 

Two alternative interpretations seem so unlikely that they can be dismissed with 
some confidence. ( 1 ) The hypothesis that the RBC and its simple Hbs had a common 
origin and that protobranch RBCs were lost in favor of a HcO 2 transport system 
after the divergence of the protobranchs and the pteriomorphs entails at least two 
independent origins of molluscan He culminating in similar quaternary structures 
with very different functional properties. As pointed out in detail earlier (Mangum el 
ai, 1985), this absence of correlation between known aspects of quaternary structure 
and respiratory function fails to provide a selection pressure for convergent evolution 
of those aspects of structure and it implies that they are conservative, ancestral charac- 
ters. The point is strengthened by the present finding of a similar quaternary structure 
of bivalve and other molluscan Hcs, with a strikingly different combination of respi- 
ratory properties. (2) The notion that the RBC had a common origin and that the 
pteriomorphs, not the protobranchs, are the more primitive bivalves can also be dis- 
missed. Abundant morphological evidence indicates otherwise. 

A third alternative, the hypothesis that the RBC had a common origin and that 
the Protobranchia and the Pteriomorpha did not have a common ancestor, is some- 
what more difficult to reject. While the possibility of a di- and even polyphyletic 
origin of the bivalves is frequently mentioned (e.g., Cox et ai, 1969; Newell, 1969; 
Runnegar, 1978), the position has been advocated positively and forcefully only with 
respect to a separate origin of the Lucinacea (McAlester, 1966), not separate origins 
of the Protobranchia and at least one other lineage containing the Pteriomorpha and 
the (infrequently) Hb-containing Heterodonta. The strongest supporting evidence 
may be the results reported by Purchon ( 1 978), who employed a matrix analysis based 
on set theory to cluster and thus to gauge the degree of relatedness between the 40 
recent bivalve superfamilies whose taxonomic integrity is uncontroversial. Using the 



BIVALVE HEMOCYANIN 219 

nine (of 1 7) characters that Newell ( 1 969) had designated as diagnostic of the super- 
families and that were either practical or suitable for the analysis, Purchon (1978) 
identified only two major clusters of bivalves, the nuculoids and the rest. His conclu- 
sion is reflected in Allen's (1986) diagnosis of the two quite different bivalve sub- 
classes. With the stipulation that the nuculoid cluster should include the solemyoids, 
as in Allen's (1986) scheme, the present findings identify a tenth character that sup- 
ports the notion of one and only one major "taxochasm" among the bivalves (Pur- 
chon, 1978). We mention the possibility of diphyly less in advocacy of it than as an 
alternative that circumvents the weaknesses of the hypothesis of multiple origins of 
the RBC and which, therefore, should not be ignored. 

Further progress in understanding the evolution of O 2 transport systems awaits 
further elucidation of the structure and function of tissue heme proteins and also 
further understanding of how bivalve O 2 transport systems work. When details such 
as blood gas levels, pH, responses to hypoxia, respiratory and cardiovascular design 
constraints, etc. are known in protobranchs as well as additional Hb-containing bi- 
valves, then the selection pressures favoring the evolution of systems with particular 
properties will become clearer. 

ACKNOWLEDGMENTS 

Supported by NSF DCB 84-14856 (Regulatory Biology), BSR 83-07714 (System- 
atic Biology), and DMB 17310 (Biochemistry). CPM is grateful to M. J. Greenberg 
for leads to the literature on bivalve phylogeny and to R. D. Barnes for helpful discus- 
sion. For MPM this is Contribution No. 153 from the Marine Science Center of 
Northeastern University. 

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Reference: Biol. Bull. 173: 222-229. (August, 1987) 



PARTICLE SIZE AND FLOW VELOCITY INDUCE AN INFERRED 
SWITCH IN BRYOZOAN SUSPENSION-FEEDING BEHAVIOR 

BETH OKAMURA 1 

Smithsonian Marine Station at Link Port, 5612 Old Dixie Highway, Fort Pierce, Florida 33450 

ABSTRACT 

The feeding rates of two bryozoan species varied with particle size and flow veloc- 
ity. In one species, increased flow reduced feeding on larger particles. The anoma- 
lously high capture rate of the largest particles by the smaller of the two species indi- 
cates a switch in feeding by ciliary currents to feeding that involves a high degree of 
tentacular activity. This is the first quantification of feeding under alternate modes 
in a benthic invertebrate and suggests that tentacular feeding may provide a signifi- 
cant source of nutrition for bryozoans. 

INTRODUCTION 

It is increasingly clear that switches in feeding strategies are common in benthic 
marine organisms. Some polychaetes, clams, and amphipods switch from deposit to 
suspension feeding with increases in flow or suspended material; some corals both 
entrap zooplankters with their tentacles and use mucus to entangle suspended par- 
ticles; and some active suspension feeders feed passively under certain conditions 
(e.g., barnacles, ascidians, brachiopods, and sponges) (Lewis and Price, 1975; Taghon 
et at, 1980; Dauer et at, 1981; LaBarbera, 1984; Olafsson, 1986). There is also evi- 
dence for alternate feeding modes in zooplanktonic suspension feeders. Copepods 
have been argued to filter feed on small particles and to actively grasp particles of 
larger size (e.g., Conover, 1966; Richman and Rogers, 1969; Poulet, 1974) [but see 
more recent clarification of copepod feeding by Koehl and Strickler (1981)]. In ad- 
dition, many suspension feeders take up dissolved organic matter, although the pro- 
cess is presumably continuous and would entail no switch in feeding behavior 
(Stephens and Schinske, 1961; JoYgensen, 1976; DeBurgh and Fankboner, 1978; 
Stewart, 1979). 

In bryozoans, feeding currents produced by cilia lining the tentacles of the lopho- 
phore can be accompanied by a high degree of tentacular activity ranging from simple 
individual tentacle-flicking to encaging particles with all of the tentacles (Winston, 
1978). Encagement activity is generally observed when particles are large; however, 
there has been no explicit test of the factors promoting greater tentacular versus ciliary 
feeding in bryozoans. In addition, the amount of food ingested under alternate modes 
has not been determined for any benthic invertebrate. This study compares the effects 
of particle size and ambient flow velocity on the feeding of the two closely related 
arborescent bryozoans, Bugula neritina and B. stolonifera. 

Bryozoans are exclusively colonial animals common in both modern and fossil 
marine habitats. A variety of colony morphologies are found among bryozoan spe- 

Received 16 March 1987; accepted 27 May 1987. 

1 Present address: Department of Oceanography, Dalhousie University, Halifax, Nova Scotia, Canada 
B3H4J1. 



222 



SWITCHING IN BRYOZOAN FEEDING MODES 223 



TABLE I 

Dimensions* of Bugula neritina andB. stolonifera 



Biigula neritina Bugula stolonifera 



Maximum size of colony (cm) 8 3-4 

Number of tentacles 23-24 13-14 

Mean length of tentacles (cm) 6.16X10" 2 4.47 X 10~ 2 

Mean diameter of lophophore (cm) 7.64 X 1CT 2 4.41 x 1(T 2 

Mean diameter of mouth (cm) 7.4X10" 3 4.9 X 10~ 3 

1 Values from Ryland and Hayward ( 1977) and Winston (1978, 1982). 



cies. These include encrusting, arborescent, and massive colonies. The zooids (mod- 
ules) that compose bryozoan colonies also vary substantially in form and function 
both among species and within colonies of the same species. Feeding zooids ingest 
suspended particles using the ciliated crown of tentacles, the lophophore. Gut con- 
tents and laboratory rearing experiments indicate that flagellates and diatoms can be 
important food items (Winston, 1977, and references therein). However, little is 
known of the food sources for field populations since plankton is inherently patchy 
in nature, the partial digestion of gut contents hinders identification of ingested mate- 
rial, and the small size of feeding zooids and the even smaller size of their prey make 
direct observation in the field difficult. The effect of flow on bryozoan feeding 
has received some attention (Okamura, 1984, 1985), but the effect of particle size 
is unknown. 

MATERIALS AND METHODS 

Colonies of the arborescent, anascan cheilostomes Bugula stolonifera and B. neri- 
tina co-occur in fouling communities that develop on submerged structures in ports 
and harbors. In Florida, colonies occur on seagrasses, oyster shells, docks, canal walls, 
floats, rotting wood, algae, coastal rock ledges, and inlet breakwaters (Winston, 1 982). 
Both species are widespread. B. stolonifera is the smaller species (see Table I) and 
often colonizes and grows within B. neritina colonies. In this study, B. neritina and 
B. stolonifera colonies were collected from the floating docks of the Harbor Branch 
Oceanographic Institution at Link Port, Florida. Laboratory observations of bryo- 
zoan feeding behavior confirm that high degrees of tentacular activity and encage- 
ment of particles occur in several Bugula species (including B. neritina) and seem to 
be associated with particle size and motility (Winston, 1978). However, these obser- 
vations were made in still water. 

Feeding experiments were performed by submerging colonies in a recirculating 
flow tank (Vogel and LaBarbera, 1978) in which currents of known mean velocities 
could be created in the working section (13 cm X 13 cm X 13 cm). The flow tank 
contained a suspension of latex particles (polystyrene divinylbenzene calibration par- 
ticles: Duke Scientific Corporation, Palo Alto, CA). (Initial observations showed that 
the bryozoans would ingest these particles.) Two flow velocities were created in the 
flow tank: a relatively slow flow (1-2 cm/s) and a relatively fast flow (10-12 cm/s). 
Flow measurements taken with an electromagnetic flow probe (Marsh McBirney No. 
523) in the field at the branch tip level of Bugula stolonifera (Okamura, 1 984) indicate 
that both species encounter flow velocities in the experimental range (measurements 



224 B. OKAMURA 

were made in a habitat where both species occurred). Feeding on three sizes of latex 
particles was assessed at each flow velocity. Particle diameters were 9.6 (SD = 0.5), 
14.6 (SD =: 1.0), and 19.1 (SD =1.1) microns. At the outset of experiments, particle 
concentrations in the flow tank were set at 100 particles/ml by adding appropriate 
volumes of stock suspensions of known concentrations to a known volume of filtered 
seawater in the flow tank. Concentrations of 100 particles/ml lie well within the range 
of concentrations of flagellates measured in the field (e.g., Jtfrgensen, 1966; Bullivant, 
1968). Control runs in the flow tank indicated that latex particles do not settle out of 
suspension over time at either flow velocity employed (Okamura, 1984). 

Up to three replicate colonies were placed in the flow tank during a given experi- 
ment. Only portions of colonies were used in all experiments. [Clipping colonies does 
not affect feeding activity (Okamura, 1984)]. Colonies were allowed to feed for 20 
min and then were removed from the flow tank and placed in dilute sodium hypo- 
chlorite. This treatment dissolves the organic contents of colonies but leaves intact 
the exoskeleton, membranous material, and latex particles. Ingested latex particles 
(that can initially be discerned only poorly in the gut before the gut wall dissolves) 
remain trapped within the zooidal exoskeleton and membranes and can be counted 
easily. These counts provided an estimate of the mean number of particles ingested 
per feeding zooid per colony (range of zooids sampled per colony = 5-100, mean 
= 64.4, SD = 35.2) (range of colonies replicated per treatment = 8-16). The effects 
of flow velocity and particle size on the mean number of particles ingested per zooid 
per colony were then analyzed with two-way analyses of variance for each species. 

RESULTS 

Bugida neritina ingested few small particles at both flow velocities (see Fig. 1A). 
More large particles were consumed than medium-sized particles in slow flow. This 
pattern reversed itself in fast flow (note the nearly significant interaction term). The 
smaller B. stolonijera showed greatest ingestion of medium-sized particles in slow 
flow (see Fig. 1 B). Feeding on medium-sized and small particles was inhibited in fast 
flow; however, large particles were captured in great numbers. 

DISCUSSION 
Feeding patterns and their causal explanations 

Rubenstein and Koehl (1977) used aerosol models of filtration to clarify suspen- 
sion-feeding mechanisms, however these models can only be applied to passive sus- 
pension feeders. Because bryozoans and other active suspension feeders generate 
feeding currents, complex three-dimensional flow patterns arise between self-gener- 
ated feeding currents and local currents near the feeding structures (Jtfrgensen, 1980). 
For feeding to occur, particles must be transferred from local currents into the self- 
generated feeding currents, and in doing so they must pass through a boundary zone 
characterized by steep velocity gradients (Jtfrgensen, 1 980). The behavior of particles 
that enter steep velocity gradients is uncertain (Strathmann, 1 982). With this in mind, 
several factors may explain the patterns of feeding from different flows on particles 
of varying size by Bugula stolonijera and B. neritina. 

Large-sized particles travel with greater momentum and thus may be carried fur- 
ther downstream before crossing flow lines in velocity gradients. Strathmann ( 197 1 , 
1982) argued that no evidence indicates that momentum carries particles across flow 
lines so that they will impinge upon the ciliary tracts of echinoderm larvae and lopho- 



SWITCHING IN BRYOZOAN FEEDING MODES 



225 



8.0- 



6.0- 



J> 2.0 
i 

TJ 

O 
O 

N 



O 



(0 

a 

6 

c 

c 

CO 
0) 

E 



1 0.0- 



8.0- 



6.0- 



4.0- 



2.0- 



neritina 




slow fast 

FLOW 




I- 
slow 



FLOW 



fast 



FIGURE I. Mean number of 9.6 (S), 14.6 (M), and 19.1 (L) micron particles captured per feeding 
zooid per colony (2 SE) by Biigula ncrilina (A) and B stolonifera (B) in slow and fast flow. Two-way 
analysis of variance for B. neritina: F, 55 (velocity term) = 1.347, P = 0.251; F 2 .5 ? (particle size term) 
= 18.120, P < 0.001; F 2 55 (velocity X particle size interaction) = 2.931, P = 0.062. Two-way analysis of 
variance for B. stolonifera: F, 59 (velocity term) = 0.035, P = 0.0851; F 2 .5 9 (particle size term) = 1 1.183, P 
< 0.00 1 ; F.,59 (velocity X particle size interaction) = 42.679, P < 0.00 1 . 



phorates, and hence that momentum does not play a role in the suspension feeding 
of these organisms. However, the role of momentum in the transport of particles out 
of local currents and into feeding currents is unknown. Alternatively, the relatively 
greater drag experienced by large-sized particles may act to sweep them further down- 
stream before crossing flow lines in velocity gradients. Both momentum and drag 
increase with ambient flow velocity. 

The larger lophophores of Bugula neritina create stronger feeding currents, and 
these may account for its greater effectiveness in capturing large particles from slow 
flow. For B. stolonifera, highest ingestion rates in slow flow were on medium-sized 
particles. In fast flow, B. neritina was hindered in feeding on large particles and 
showed highest ingestion rates on intermediate-sized particles. The greater momen- 
tum of or drag on large particles in faster flow may make their ingestion more difficult. 
However, anomalously high feeding rates on large particles in fast flow were observed 
for B. stolonifera. The most likely explanation for this is a switch in feeding from 
mainly ciliary currents to feeding that involves a high degree of tentacular activity. 
Unfortunately, a switch in feeding technique was not anticipated and, consequently, 
lophophore behavior was not observed with a microscope during the experiments. 



226 B. OKAMURA 

Furthermore, colonies were fixed and sampled when time permitted. When the feed- 
ing patterns were eventually discerned, there were no colonies available for observing 
lophophoral behavior (both species are highly seasonal in Florida). However, as men- 
tioned earlier, laboratory observations indicate that feeding in many Bugula species 
does involve high degrees of tentacular activity and encagement (Winston, 1978). 
Only moderate levels of tentacular activity were observed in B. stolonifera, but since 
Winston's observations were made in still water and the sizes of suspended particles 
were unspecified, conditions that would have invoked high degrees of tentacular ac- 
tivity or encagement by B. stolonifera may not have been present. 

The inferred switch in feeding mode by Bugula stolonifera is induced by increased 
flow rate and depends on particle size. The larger B. neritina showed no evidence of 
a switch in feeding behavior. However, if particles of larger size or perhaps if faster 
flow velocities had been employed, a switch in feeding would be expected. High de- 
grees of tentacular activity were observed for B. neritina, including the formation of 
cages with its tentacles (Winston, 1978). The reduction in feeding on small and medi- 
um-sized particles with increased flow by B. stolonifera is in accord with previous 
results (Okamura, 1985). 

Best and Thorpe ( 1 983, 1 986) provide evidence that bryozoans are capable of altering 
the strength of their feeding currents and do so in response to particle concentration. An 
alternate explanation for the present results is that the feeding patterns are produced by 
feeding currents of different strengths. If this is so, the greater flux of large particles in fast 
flow would induce Bugula stolonifera, but not B. neritina, to produce stronger feeding 
currents. While this is a possibility, it is considered unlikely since B. stolonifera fed dispro- 
portionately on large particles present in mixtures (composed of equal proportions of all 
three particle sizes) in fast flow (Okamura, in prep). In this case, if stronger feeding cur- 
rents were produced, particles of all three sizes would be expected to be ingested in equal 
proportions. It is more likely that the disproportionate ingestion of large particles from 
mixtures was a result of selective tentacular feeding. 

It is notable that the apparent switch in feeding by Bugula stolonifera entails such 
a large increase in capture. Tentacular feeding may involve a much greater energetic 
expenditure than ciliary feeding. Only when the gain is great (i.e., many large particles 
per unit time) will feeding that involves a high degree of tentacular activity be a worth- 
while strategy. Note that a switch to tentacular feeding results in a much greater 
amount of "biomass" captured [mean mass of large particles captured by B. stoloni- 
fera in fast flow - 30.80 X 19~ 9 g (SD == 8.50), of medium-sized particles = 6.21 
X 10" 9 g(SD= 1.87), and of small particles = 1.09X 10~ 9 g(SD = 0.42)]. However, 
the costs of ciliary and tentacular feeding are unknown. The reduced surface area 
offered by small particles for tentacular contact may preclude tentacular feeding on 
particles below a minimum size, or perhaps particles must exceed a certain relative 
size to be perceived individually. 

Plasticity in feeding behavior and its implications 

It is evident that many suspension feeders are capable of great plasticity in feeding 
behavior. Alternate feeding techniques are invoked by variations in the suspension 
from which they capture their food. These variations may be characteristics of the 
prey items (e.g., size, motility, chemistry) or physical properties of the medium itself 
(e.g., temperature, density, and the patterns of fluid flow). Since suspension feeders 
will regularly encounter suspensions that vary in both physical properties and prey 
items, plasticity in feeding response is expected. The study of suspension feeding in 



SWITCHING IN BRYOZOAN FEEDING MODES 227 

still water on uniform particles may often provide an incomplete picture of the feed- 
ing of many organisms. This is exemplified by the studies of Best and Thorpe (1983, 
1986). They argue that tentacular flicking and the more localized ciliary reversal 
mechanism that Strathmann (1982) proposes to account for particle capture during 
ciliary feeding are not the main methods of feeding employed by bryozoans. They 
suggest that, overall, the bulk of particles ingested are those that feeding currents carry 
down the center of the lophophore towards the mouth. The importance of ciliary 
reversal and tentacular activity in feeding are rejected on the basis of calculating the 
number of reversals and tentacular flicks required to explain the ingestion rates they 
observed. However, their evidence may be biased due to their use of extremely high 
particle concentrations (50-200 cells M! '), small particle sizes, and the absence of 
ambient currents in their experiments. Their study suggests that very high particle 
concentrations may swamp contributions to feeding by mechanisms other than the 
bulk flow of particles through the center of the lophophore, while this study suggests 
that high degrees of tentacular activity depend on both particle size and ambient flow. 

Many investigators have studied the relationship between bryozoan colony form 
and the patterns of self-generated feeding currents through colonies (Cowen and 
Rider, 1972; McKinney, 1977, 1986a, b; Taylor, 1979; Anstey, 1981; McKinney et 
ai, 1986). Results reported here indicate that feeding currents may not always be of 
primary importance in particle capture. The potential for alternate feeding behaviors 
should be appreciated when interpreting colony morphology solely in terms of feed- 
ing current patterns. Both stenolaemate and gymnolaemate bryozoans display high 
degrees of tentacular activity even in still water (Winston, 1978). Tentacular feeding 
from faster ambient flow and/or on certain types of prey may provide a significant 
source of nutrition for a variety of bryozoans. 

Optimal foraging theory attempts to explain and predict many aspects of the for- 
aging behavior of animals by assessing foraging tactics in terms of maximizing net 
rates of energy gain and therefore fitness (e.g., Schoener, 1971; Pyke 1977, 1984; 
Hughes, 1980). Which prey will be the "best" is determined by the energy content of 
the prey and the energetic cost to the predator of searching for and handling the 
prey. Thus, understanding patterns of prey selection, prey vulnerability, and feeding 
behaviors is crucial in the interpretation of foraging strategies. Particle size appears 
to relate to prey vulnerability in bryozoan feeding. Flow velocity induces a switch in 
feeding behavior that results in a shift in the size of particles captured. Furthermore, 
flow velocity appears to control the vulnerability of particles of certain size ranges 
even when feeding under one mode (note greater feeding on large particles in slow 
flow but on medium-sized particles in fast flow by Bugula neritind). Prey vulnerabil- 
ity and patterns of prey capture are thus determined by both the constraints imposed 
by flow and by the flow-induced change in feeding tactics. This suggests that the role 
of flow on particle size selection and the behavior of suspension feeders merits further 
investigation. In addition, a switch in feeding tactics by bryozoans implies that these 
organisms perceive and assess prey availability and subsequently adopt the most 
efficient feeding mode (i.e., the one that maximizes net energy gain). It appears that 
predictions of optimal foraging theory may be applicable to benthic suspension feed- 
ers despite their seemingly simple sensory capabilities and sessile existence. 

ACKNOWLEDGMENTS 

I thank A. H. Cheetham, M. A. R. Koehl, F. K. McKinney, M. E. Rice, P. D. 
Taylor, and two referees for suggesting improvements to the manuscript. This is con- 
tribution #1 77 of the Smithsonian Marine Station at Link Port. 



228 B. OKAMURA 

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Reference: Biol. Bull. 173: 230-238. (August, 1987) 



EPITHELIAL WATER PERMEABILITY IN THE EURYH ALINE MUSSEL 

GEUKEi 'SIA DEMISSA: DECREASE IN RESPONSE TO HYPOOSMOTIC 

MEDIA AND HORMONAL MODULATION 1 

LEWIS E. DEATON 2 

Whitney Marine Laboratory, University oj Florida, Route I, Box 121, St. Augustine, Florida 32084 

ABSTRACT 

The diffusional water permeability of isolated mantles from the mussel Geukensia 
demissa was reduced by incubation of the tissues in hypoosmotic media. The perme- 
ability of mantles from 1000 mOsm seawater (SW)-accli mated animals was 6 X 10"' 
cm/s. A four-hour incubation in 500 mOsm SW or 250 mOsm SW reduced the water 
permeability by 2 X 10~ 5 cm/s and 4 X 10 5 cm/s, respectively. A half-hour exposure 
to the hypoosmotic medium was sufficient to induce the decrease in permeability. 

The water permeability of mantles incubated in isosmotic SW containing acetone 
extracts of ganglia from 1000 mOsm SW-acclimated mussels or of mantle from 500 
mOsm SW-acclimated mussels was significantly reduced. Extracts of gill had no 
effect. 

Ovine prolactin (50 mg/ml) decreased the water permeability of mantles in isos- 
motic seawater. Cortisol ( 10~ 4 M), arginine vasopressin ( 10~ 6 M), and the molluscan 
neuropeptide FMRFamide ( 10 6 M) had no effect. 

These results show that the epithelial water permeability of euryhaline bivalves 
varies with changes in the ambient salinity, and that these permeability changes may 
be modulated by factors of neural origin. 

INTRODUCTION 

A number of specific physiological mechanisms facilitate the survival of euryha- 
line marine animals in habitats characterized by variations in salinity. These mecha- 
nisms include changes in urinary output, regulation of the extracellular fluid compo- 
sition, volume regulation, and changes in epithelial permeability to water and ions. 

A reduced epithelial permeability to water in dilute media has been reported in 
several invertebrates, including a number of arthropods (Rudy, 1967; Smith, 1970a; 
Capen, 1972; Smith and Rudy, 1972; Cornell, 1973; Hannen and Evans, 1973; Lock- 
wood et ai, 1973; Roseijadi et ai, 1976; Thuet, 1978), three polychaetes (Smith, 
1964, 1970b; Fletcher, 1974), and a bivalve (Prusch and Hall, 1978). The phenome- 
non has been observed in osmoconformers (e.g., Mytilus edulis, Libinia emarginatd) 
and in osmoregulators (e.g., Rhithropanopeus harrisi, Nereis limnicola). Nearly all of 
these data were collected by measuring changes in the fluxes of water into or out of 
whole animals exposed to dilute media. As indicated by Cornell (1979), part of the 
observed decreases in the flux of water across the epithelium of an intact animal could 
be effected by changes in circulation or ventilation. This criticism does not apply to 



Received 9 February 1987; accepted 26 May 1987. 

' This is contribution number 265 from the Tallahassee, Sopchoppy and Gulf Coast Marine Biological 
Association. 

2 Current address: Department of Biology, University of Southwestern Louisiana, Lafayette, Louisi- 
ana 70504. 

230 



EPITHELIAL WATER PERMEABILITY IN G. DEMISSA 231 

studies of the response of the epithelial permeability of isolated tissues to changes in 
salinity. 

Isolated tissues from only a few marine invertebrates have been used to examine 
changes in water permeability in response to dilution of the medium. Cantelmo 
(1977) found that the water permeability of gut epithelia and gills from the crabs 
Cancer irroratus and Callinectes sapidus was lower in tissues isolated from animals 
acclimated to 40% seawater than in tissues from animals acclimated to 100% seawa- 
ter. However, exposure of isolated tissues to hypoosmotic media for 2 h did not 
change their water permeability. Acclimation of the mussel Mytilus edulis to reduced 
salinity caused a decrease in the diffusional permeability of water across isolated man- 
tle tissues (Prusch and Hall, 1978). 

Neurohormones have been implicated in the modulation of water permeability 
in a variety of invertebrates. For example, extracts of the thoracic ganglion decrease 
the water flux across isolated crab gut and gills (Mantel, 1968; Berlind and Kamem- 
oto, 1977), and the injection of a brain homogenate reduces the rate of water ex- 
change of earthworms (Carely, 198 1 ). Two lines of evidence suggest a possible modu- 
lation of water balance by neural products in gastropod molluscs. Injection of syn- 
thetic thyrotropin releasing hormone into the freshwater snail Lymnaea stagnalis 
causes a slight loss in wet weight (Grimm-Jorgenson, 1979). Similarly, injection of 
homogenized R-15 cells from the abdominal ganglion of the opisthobranch Aplysia 
brasiliensis resulted in a 5% gain in wet weight (Kupfermann and Weiss, 1976). 

The present study was undertaken to determine the response of the epithelial 
water permeability of isolated mantle tissue from the euryhaline mussel Geukensia 
demissa to decreases in the ambient osmotic concentration. This tissue was also used 
as a bioassay system to determine the effects of extracts of the ganglia and other tissues 
on water permeability. 

The results show that the diffusional water permeability (P d ) of the mussel de- 
creases in response to the dilution of the external medium, and that water permeabil- 
ity may be modulated by a factor of neural origin. 

MATERIALS AND METHODS 
Animals 

Atlantic ribbed mussels, Geukensia demissa granosissima, were collected from a 
salt marsh near St. Augustine Beach, Florida. The animals were kept unfed in running 
seawater (30%o) at ambient temperature. All animals were used within 3 weeks of 
collection. 

Histology 

The adductor muscles of individual mussels were cut, the animals opened, and 
pieces of the central portion of the mantle dissected free. The tissue was fixed in 
filtered seawater containing 2% glutaraldehyde, dehydrated and cleared in a graded 
series of water/ethanol/t-butyl alcohol, and embedded in paraffin. Sections (10 ^m) 
were cut on a microtome, stained with hematoxylin eosin, and mounted on glass 
slides. 

Measurement of diffusional water permeability 

The adductor muscles of individual mussels were cut and the valves carefully 
pried apart. Each mussel provided two tissues, the mantle covering the inside of the 



232 L. E. DEATON 

right valve and the mantle covering the inside of the left valve. The left and right 
mantle halves were cut away from the visceral mass and detached from the margins 
of their respective valves. The isolated tissues were placed in small covered dishes 
containing 5 ml of 1000 mOsm seawater (SW) which was aerated via small bore 
tubing. After 60 minutes of incubation, one mantle half was mounted over the aper- 
ture of one half of a diffusion chamber and secured with a soft rubber o-ring. The 

jes were always mounted in the chamber so that the movement of tritated water 
was from the extrapallial cavity side to the mantle cavity side. Five ml of medium 
were placed in the chamber to provide a hydrostatic pressure head sufficient to check 
the mounted tissue for obvious leaks. The chamber was then assembled and both 
sides filled with medium by alternate additions of 2-3 mis. The total volume of each 
compartment of the chamber was 1 4 ml; the area of exposed tissue was 1 cm 2 . Mixing 
and aeration were provided by gas lift pumps powered by water-saturated air. About 
1 /uCi of tritiated water was added to one compartment of the chamber (the "hot" 
side), and following 10 minutes equilibration, 100 yul samples of the medium in the 
other compartment (cold side) were removed at 1 5 minute intervals for 60-90 min- 
utes. These samples were mixed with 10 ml scintillation cocktail and counted on a 
liquid scintillation counter. In experiments using paired left and right mantles, the 
flux across one mantle was measured while the matching tissue was transferred to a 
dish containing either control (1000 mOsm SW) or experimental medium. These 
matching tissues were further incubated from one half to four hours and then their 
P d values determined in the same rinsed and dried chamber used for the matching 
control measurement. 

The water flux across the tissue and the diffusional water permeability were calcu- 
lated from equations 1 and 2, respectively: 

/ 1 \ Q* / 1 \ Q*/Qi = specific activity in compartment 1 
1- Ji2 2 = I " I ~ I A I Q? = amount of isotope in compartment 2 

VUQf/Q, W - 



-time 

J P H 2 A = area of tissue 

*a C w = molar concentration of water 



The differences in P d values between paired left and right mantles were averaged 
and differences among treatment means assessed by Student's / test. 

Tissue extracts 

The pedal, visceral, and cerebral ganglia from 250 mussels acclimated to 1000 
mOsm SW were dissected from the animals and pooled in a large volume of cold 
acetone to extract putative hormones and inactivate proteolytic enzymes; acetone 
extracts were also made of the gills and mantles from these mussels. The mantles of 
200 mussels acclimated to 500 mOsm SW were also extracted in acetone. The extracts 
were evaporated to dryness on a rotary evaporator and the water soluble portion of 
the residue taken up in a minimal volume of distilled water. The dose added to the 
incubation media was approximately 1 animal equivalent. All tissue extracts and 
hormones were added to the incubation media and to the fluid in both compartments 
of the diffusion chamber. 

RESULTS 
Histology 

Figure 1 shows a cross section of the central portion of the mantle. This complex 
tissue separates the extrapallial space from the mantle cavity. Both surfaces are lined 



EPITHELIAL WATER PERMEABILITY IN G. DEMISSA 



233 



MC 




EPS 



FIGURE 1. Cross-section through the central portion of the mantle ofGenkensia demissa. A. Section 
from the mantle cavity (MC) to the extrapallial space (EPS) showing the epithelia (epi) on both surfaces 
with underlying muscle layers (m). The epithelia are separated by connective tissue which encompasses 
many hemolymph vessels (*), few genital follicles or canals (g), and transverse muscle bundles (tmb) con- 
necting the two subepithelial muscle layers. Bar = 200 ^m. B. Columnar extrapallial epithelium with thin 
underlying muscle layer. Bar = 100 ^m. C. Squamous mantle cavity epithelium with thick underlying 
muscle layer. Fibers from the transverse muscle bundles splay out to join the subepithelial muscle layer 
(arrows in C2). Bar = 100 



with an epithelium underlain by a muscle layer (Fig. 1 A). The extrapallial epithelial 
cells are much taller than those lining the mantle cavity, but the subepithelial muscle 
layer associated with the extrapallial space is much thinner than that on the mantle 
cavity side of the tissue (compare Figs. 1 A, ICi). The bulk of the mantle is occupied 
by connective tissue in which are found numerous hemolymph vessels and, in these 
non-reproductive specimens, occasional genital canals (Fig. 1A). Bundles of muscle 
fibers traverse the mantle joining the two subepithelial muscle layers (Figs. 1A, 
1C^). Similar structures have been described for the mantles of other species (Beed- 
ham, 1958). 

Diffusional water permeability 

Preliminary experiments showed that the accumulation of counts in the cold 
compartment of the diffusion chamber was linear with time for over six hours, indi- 
cating that the 10 minute equilibration with labelled water was sufficient for attain- 
ment of a steady-state flux across the tissue. The diffusional water permeabilities of 



234 L. E. DEATON 

TABLE I 

Changes in diffisional water permeability (P d ) of mantles from 1000 mOsm seawater-acclimated 
Geukensia demissa after a four hour incubation in various seawaters 

Treatment medium lOOOmOsm 500 mOsm 250 mOsm 
Change in P d 0.5 x 10 5 ***-2.3 X 10~ 5 ***_ 4 .! x 10 -s 
SD 1.1X10' 5 3.0X10 5 1.3X10' 5 
n_ ^8 9 7 

Values are in cm/s and represent differences in P d between paired left and right mantles: one mantle 
of each pair was incubated in 1000 mOsm SW for 1 h; the other was incubated in the treatment medium 
for 4 h. Values significantly different from the 1000 mOsm treatment are marked with *** (P < .00 1 ). 



mantles from animals acclimated to 1000 and 500 mOsm for three weeks were, re- 
spectively, 7.9 3.3 X 10" 5 (n = 10) and 4.3 0.7 X 10 5 cm/s). These values are 
higher than the mean (2.2 X 10" 5 cm/s obtained by Prusch and Hall (1978) for man- 
tles of G. demissa acclimated to 1000 mOsm SW. Their animals, collected near 
Woods Hole, Massachusetts, were undoubtedly the subspecies G. demissa demissa. 
Differences in chamber design and differences between the two G. demissa subspecies 
probably account for the discrepancy. 

There were no significant differences between the mean P d values of mantles incu- 
bated for four hours in 1000 mOsm SW (6. 1 2.9 X 10" 5 cm/s) and the mean P d s of 
the matching control (incubated in 1 000 mOsm S W for 1 h) tissues (6.5 2.9 X 10~ f 
cm/s), nor was the mean of the differences between paired tissues (0.5 1.1 X 10~ f 
cm/s) significantly different from zero. In contrast, the water permeability of mantles 
incubated in 500 or 250 mOsm SW for four hours was decreased by '/3 and %, respec- 
tively, compared to paired controls (Table I). The data from a representative experi- 
ment are shown in Figure 2. The movement of labelled water across both tissues is 
linear with time; the movement of water across the tissue incubated in 500 mOsm 
SW is slower. The magnitude of the reduction in flux is constant throughout the 
experiment. 

The time course of the reduction of epithelial water permeability (P d ) in 500 
mOsm SW is summarized in Table II. A thirty minute incubation in dilute seawater 
was sufficient to induce a decrease of about 1 X 10~ 5 cm/s in the P d value. Longer 
incubations further reduce the permeability, but these values were not significantly 
different from that induced by a thirty minute incubation (Table II). 

When mantles were incubated in isosmotic medium containing an extract of gan- 
glia from 1000 mOsm SW-acclimated mussels or an extract of mantles from 500 
mOsm SW-acclimated mussels, the water permeabilities were significantly reduced. 
Extracts of other tissues had no effect (Table III). 

Hormones which affect the water permeability of vertebrate tissues were tested 
for effects on the bivalve mantle. Ovine prolactin significantly reduced the P d value 
of mantle tissues in isosmotic media. Neither arginine vasopressin nor cortisol 
changed the permeability of mantles in isosmotic SW (Table IV). The small reduction 
in the P d value of tissues incubated in isosmotic media with the molluscan neuropep- 
tide FMRFamide was not significant (Table IV). Colchicine did not prevent the de- 
crease in P d induced by exposure to dilute media (Table V). 

DISCUSSION 

The diffusional water permeability (P d ) of isolated mantles from the euryhaline 
mussel Geukensia demissa decreases when the tissues are exposed to hypoosmotic 



EPITHELIAL WATER PERMEABILITY IN G. DEM1SSA 



235 



30 



25- 



20 



c pm 
xlO 3 l5 



10 




.0 



15 30 45 

Time (min) 



60 



FIGURE 2. The unidirectional movement of Initiated water across a piece of isolated mantle ofGeuken- 
sia demissa. Total counts per minute appearing in the "cold" side of a diffusion chamber are plotted as a 
function of time. The data are from paired mantle tissues from one mussel: the left mantle was incubated 
for 1 h in 1000 mOsm seawater (solid circles); the right mantle was incubated in 500 mOsm seawater (open 
circles) prior to measurement of the tritiated water flux in a diffusion chamber containing the same media. 



media; the decrease in permeability is proportional to the magnitude of the decrease 
in the ambient osmotic concentration. Furthermore, the P d of the isolated tissue incu- 
bated in isosmotic medium is reduced by a vertebrate hormone and by an endoge- 
nous factor of neural origin. 

The reduction in permeability induced by a 30 min exposure to 500 mOsm seawa- 
ter is less than that resulting from long-term acclimation of the mussels to 500 mOsm 
SW. The reduction of water permeability by ganglion extracts from 1000 mOsm-SW 
acclimated mussels and mantle extracts from 500 mOsm-SW acclimated mussels 
suggests that the putative factor is produced in the ganglia and released to the periph- 



TABLE II 

Time course of change in water permeability (PJ of mantles from WOO mOsm seawater-acclimated 
Geukensia demissa during incubation in 1000 mOsm or 500 mOsm seawater 

Treatment medium 



lOOOmOsm 



500 mOsm 



Incubation duration (h) 


0.5 


1 


2 


3 


4 


0.5 


1 




2 


3 


4 


P d change (XlO~ 5 cm/s) 


0.9 


0.4 


0.3 


0.4 


0.2 


*-0.9 


***-!. 





**-1.4 


***-!. 9 


***-!. 9 


SD 


1.1 


0.4 


1.4 


0.9 


0.9 


0.6 


0. 


9 


1.4 


1.8 


1.4 


n 


4 


3 


4 


8 


5 


4 


10 




9 


10 


7 



Values are differences in P d between paired left and right mantles: one mantle was incubated in 1000 
mOsm SW for 1 h; the other was incubated in the treatment medium for 0.5 to 4 h. Values marked with 
asterisks are significantly different from the corresponding 1000 mOsm treatment value (* = P < .05, ** 
= /><. 01, *** = /><. 001) 



236 L. E. DEATON 

TABLE III 

The effect of various tissue extracts on the water permeability (PJ of mantles from WOO mOsm seawater 
acclimated Geukensia demissa incubated four hours in 1000 mOsm seawater 

Treatment medium 







1000SW 


1000SW 


1000SW 


1000SW 






+ 1000 


+ 1000 


+ 1000 


+ 500 




1000SW 


mantle ext. 


gill ext. 


ganglia ext. 


mantle ext. 


P d change 


0.5 


1.3 


0.6 


*-1.4 


**-1.5 


SD 


1.4 


1.1 


1.6 


1.2 


1.9 


n 


18 


6 


5 


3 


10 



Values are xlO 5 cm/s and represent differences in P d between paired left and right mantles: one 
mantle was incubated in 1000 mOsm SW for 1 h; the other was incubated in the treatment medium for 4 
h. Values marked by asterisks are significantly different from that for 1000SW(* = P< .05;** = P< .01). 



ery during acclimation to low ambient salinity. Prusch and Hall (1978) observed a 
67% reduction in the water permeability of mantles isolated from the mussel Mytilus 
edulis during four weeks of acclimation to 70% seawater. Thus, in the intact animal, 
continuous release of neural factors may facilitate a larger decline in permeability 
than occurs in isolated tissues. As yet there are no data on the size, structure, or 
chemical nature of this putative neurohormone. However, it is not FMRFamide (Ta- 
ble III). 

Prolactin reduced the water permeability of G. demissa mantles in isosmotic me- 
dia (Table IV). Prolactin also reduces the permeability of teleost epithelia to water 
and ions (Doneen and Bern, 1974; Foskett et al, 1983). While the presence of prolac- 
tin has been demonstrated by immunocytochemical methods in ascidians (Pestarino, 
1984), it has not been reported in any other invertebrate. Therefore it is unlikely that 
the active substance in G. demissa ganglia is prolactin. 

Khan and Salueddin (1979; 198 1 ) associated changes in the anatomy of the sep- 
tate junctions between kidney cells in the snail Helisoma duryi with increased water 
permeability. Extracts of the visceral ganglia induce these changes which occur within 



TABLE IV 

The effects of selected hormones on the change in diffusional water permeability (PJ of mantles from WOO 
mOsm seawater-acclimated Geukensia demissa incubated four hours in WOO mOsm seawater 

Treatment medium 













1000SW 






1000SW 


1000SW 


1000SW 


+ Arg 






+ Prolactin 


+ FMRFamide 


+ cortisol 


vasopressm 




1000SW 


(50 M g/ml) 


(\0~ 6 M) 


( 10~ 4 M) 


(10- 6 M) 


P d change 


0.5 


***-2.3 


-0.4 


-0.5 


-0.4 


SD 


1.1 


1.4 


0.9 


1.5 


2.0 


n 


18 


6 


4 


4 


5 



Values are XlO 5 cm/s and represent differences in P d between paired left and right mantles: one 
mantle was incubated in 1000 mOsm SW for 1 h; the other was incubated in the treatment medium for 4 
h. The value marked by *** is significantly different from that for 1000 SW (P < .001). 



EPITHELIAL WATER PERMEABILITY IN G. DEMISSA 237 

TABLE V 

The effect of colchicine on the water permeability (Pd) of mantles from 1000 mOsm seawater-acclimated 
Geukensia demissa incubated four hours in 500 mOsm seawater 

Treatment medium 



500 SW + colchicine 
500 SW (2X10~ 4 A/) 



P d change -2.3 -2.0 

SD 3.0 3. 1 

n 9 3 

Values are X 10~ 5 cm/s and represent differences in P d between paired left and right mantles: one mantle 
was incubated in 1000 mOsm SW for 1 h; the other was incubated in the treatment medium for 4 h. 

30 minutes. Neurohormones, then, can alter the water permeability of molluscan 
epithelia by causing changes in the structure of the tissues, thereby changing the resis- 
tance of the paracellular pathway to water movement. The failure of colchicine to 
prevent a decrease in the P d of mantles exposed to dilute media (Table V) suggests 
that microfilament activity is not involved in the process. 

If the major route of water movement across the G. demissa mantle is paracellu- 
lar, osmotic swelling of the epithelial cells could contribute to a decrease in the water 
permeability of the tissue during exposure to hypoosmotic media. Isolated G. demissa 
ventricles exposed to hypoosmotic seawater stop beating, but the mechanical activity 
of the ventricle recovers within 90-120 minutes (Pierce and Greenberg, 1972). Re- 
covery of the mechanical activity of the ventricle apparently is due to cellular volume 
regulation. If the time course of recovery of cellular volume by the mantle cells is 
similar to that of the myocardial cells, osmotic swelling cannot account for the reduc- 
tion in water permeability induced by 2-4 h incubations in hypoosmotic seawater. 
However, in the absence of data on the time course of changes in the volume of 
mantle cells exposed to hypoosmotic stress, the possibility that cell swelling accounts 
for some or all of the decrease in epithelial permeability cannot be ruled out. 

The mantle of G. demissa is vascularized (Fig. 1 ) and therefore well-perfused by 
the circulation. Mounting the tissues in the diffusion chamber precluded perfusion, 
and therefore the effects of delivery of the tissue extracts and other drugs via the 
circulation cannot be assessed. 

Extracts of various nervous tissues affect epithelial water permeability of crusta- 
ceans and annelids (Mantel, 1968; Tullis and Kamemoto, 1974; Berlind and Ka- 
memoto, 1977; Carely, 1981). While none of these factors has yet been identified, it 
is clear that neural factors modulate water permeability in euryhaline invertebrates. 

In summary, the epithelial water permeability of euryhaline molluscs changes 
during acclimation to changes in the ambient salinity. These changes in permeability 
may be modulated by one or more neural factors of unknown structure. The mecha- 
nisms responsible for increases or decreases in water permeability apparently involve 
changes in the junctional complexes between the epithelial cells, but factors affecting 
transcellular water permeability, such as the insertion or removal of water channels 
or changes in the composition of the membrane lipid bilayer, cannot be ruled out. 

ACKNOWLEDGMENTS 

I thank Dr. Michael J. Greenberg for critical readings of the manuscript. This 
work was supported by a grant (PCM 83093 14) to MJG and LED from the National 
Science Foundation. 



238 L. E. DEATON 

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and in isolated perfused gills. Comp. Biochem. Physiol. 58A: 382-385. 
CANTELV >, A. C. 1977. Water permeability of isolated tissues from decapod crustaceans. 1. Effect of 

iolic conditions. Comp. Biochem. Physiol. 58A: 343-348. 
CAPEN. R. L. 1972. Studies of water uptake in the euryhaline crab Rhithropanopeus harrisi. J. Exp. Zool. 

182:307-319. 
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8: 433-442. 
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Reference: Biol. Bull. 173: 239-251. (August, 1987) 



METAL REGULATION AND MOLTING IN THE BLUE CRAB, 
CALLINECTES SAPIDUS: METALLOTHIONEIN FUNCTION 

IN METAL METABOLISM 

DAVID W. ENGEL 1 AND MARIUS BROUWER 2 

' National Marine Fisheries Service, NOAA, Southeast Fisheries Center, Beaufort Laboratory, Beaufort, 
North Carolina 28516-9722 and 2 Duke University Marine Laboratory, Marine Biomedical Center, 

Beaufort, North Carolina 28516-9722 

ABSTRACT 

We recently demonstrated that zinc, copper, and hemocyanin metabolism in the 
blue crab varies as a function of the molt cycle. To extend these observations, and 
better delineate metal metabolism in marine crustaceans, we have conducted experi- 
ments to determine if environmental temperature and season of the year affect con- 
centrations of hemocyanin and copper in the hemolymph and copper and zinc in the 
digestive gland. Overwintering, cold water crabs (6C) had decreased hemocyanin 
and copper in the hemolymph and normal zinc and copper in the digestive gland 
with respect to summer crabs collected at 20-30C. When these crabs were warmed 
to 20C and fed fish for three weeks, they showed increases in the concentrations of 
copper in the digestive gland, and copper and hemocyanin in the hemolymph. In 
addition, a change from a zinc to a copper-dominated metallothionein was found in 
a majority of the warmed crabs, suggesting the involvement of copper metallothio- 
nein in the resynthesis of hemocyanin. Based on these observations and previous data 
(Engel, 1987) a conceptual model of copper and zinc partitioning in the blue crab 
has been constructed. In this model, metallothionein has an important role in metal 
regulation both during molting and in the changes related to season of the year. Met- 
allothionein-bound copper and zinc appear to be regulated at the cellular level for 
the synthesis of metalloproteins, such as hemocyanin (copper) and carbonic anhy- 
drase (zinc), both of which are necessary for normal growth and survival. Finally, we 
present evidence showing that copper metallothionein can directly transfer its metal 
to the active site of apohemocyanin. Copper insertion seems to precede the formation 
of viable oxygen binding sites. 

INTRODUCTION 

Studies of the reputed function of metallothionein in marine organisms have been 
concerned primarily with its role in detoxifying elevated concentrations of trace-met- 
als accumulated from polluted environments (Roesijadi, 1981; Engel and Brouwer, 
1984; and George and Viarengo, 1985). These reviews discussed the potential consti- 
tutive or regulatory function of metallothionein in metal metabolism, but empha- 
sized its role in detoxifying metals. 

Previously we alluded to the possibility that metallothioneins may play a role in 
organismal and cellular metal metabolism in marine species (Engel and Brouwer, 

Received 9 February 1987; accepted 19 May 1987. 



239 



240 D. W. ENGEL AND M. BROUWER 

1984; Engel and Roesijadi, 1987). Such suggestions also have been made concerning 
zinc and copper metabolism in mammals. Cousins (1982, 1985) discussed the role 
of metallothionein in zinc metabolism in rats. The observation that glucocorticoid 
hormones can significantly alter zinc metabolism and increase metallothionein syn- 
thesis in the liver without the administration of exogenous zinc also supports the 
hypothesis that metallothionein is active in normal metal metabolism (Karin, 1985). 
More recently Petering and Fowler (1986) discussed the normal or constitutive as- 
pects of metallothionein synthesis and turnover in mammals, and correlations also 
were made with non-mammalian organisms. There is a growing body of evidence, 
therefore, that metallothioneins are indeed involved in the regulation of metal me- 
tabolism. 

Recently it was demonstrated that blue crabs collected from unpolluted environ- 
ments significantly alter tissue metal concentrations and the metal composition of 
metallothionein during the molt cycle (Engel, 1987). These studies clearly demon- 
strated that metallothionein in a marine crustacean is actively involved in normal 
physiological and biochemical processes of metal regulation at the cellular level that 
control growth and reproduction. Additionally, blue crab metallothionein also is as- 
sociated with cellular metal detoxification and sequestration (Brouwer et ai, 1984; 
Engel and Brouwer, 1984). 

Two series of experiments were performed to explore further the role of metallo- 
thionein in metal metabolism. The first series of experiments examined the effect of 
overwintering on the metal metabolism of the blue crab at both the tissue and cyto- 
solic level. The second series of experiments measured the ability of metallothionein 
to donate copper for activation of apohemocyanin in vitro. In addition, we discuss 
how metallothionein and the metals bound to it relate to the physiological and bio- 
chemical changes that occur during molting. We also propose a model for the direct 
involvement of metallothionein as a metal donor in the synthesis of hemocyanin and 
zinc enzymes. 

MATERIALS AND METHODS 

All crabs used in these experiments were captured in the vicinity of Beaufort, 
North Carolina, by commercial fishermen. A group often intermolt (C 4 ) male blue 
crabs (Callinectes sapidus) were obtained in February 1986, and were maintained 
in the laboratory at ambient temperature and salinity (6C, 30%o). After a week of 
acclimation, five crabs were taken for hemolymph and tissue samples. The remaining 
five crabs were held for an additional three weeks, and water temperature was allowed 
to increase to about 20C in 10 days. During the three week period the crabs were fed 
chopped fish every other day. At the end of three weeks the remaining five crabs were 
killed and hemolymph and digestive gland samples were taken. 

Tissue metal measurements 

The concentrations of copper and zinc were determined in samples of digestive 
gland and hemolymph from individual blue crabs. The hemolymph samples were 
collected by severing the fifth pereiopod at the meropodite and collecting the fluid in 
a polyethylene vial. A portion of the hemolymph was taken for metal analysis and 
the remainder was used for determination of hemocyanin concentration. The crabs 
were killed by removing the carapace, and the digestive gland was dissected out and 
used for total metal measurements and cytosolic metal determinations. The tissue 



BLUE CRAB METAL METABOLISM 



241 



TABLE I 
Amino acid composition of blue crab and lobster metallothionein (Residues/6500 Daltons) 

Blue crab 



Lobster 





CdMT a gill 


CdMT h 
digestive gland 


ZnMT b 
digestive gland 


CuMT c 
digestive gland 


Cysteine 


18 


17 


18 


18 


Asp/Asn 


4 


4 


4 


3 


Thr 


3 


5 


5 


4 


Ser 


7 


5 


6 


5 


Glu/Gln 


8 


7 


6 


4 


Pro 


5 


6 


5 


6 


Gly 


5 


7 


6 


5 


Ala 


2 


3 


3 


3 


Val 


1 


1 


1 





Met 














He 














Leu 





1 


1 





Tyr 














Phe 














His 


1 











Lys 


7 


7 


7 


8 


Arg 


1 


1 


1 


1 




62 


64 


63 


57 



3 Brouwer el at. 1 984. 
h Brouwer unpub. results. 
c Brouwer el al 1986. 



that was used for determination of cytosolic distribution of metals was frozen rapidly 
and stored in a freezer at -70C. 

Tissue samples used for metal analysis were oven dried at 100C for 48 h and wet 
ashed with concentrated HNO 3 at 90C. Residue was dissolved in 0.25 N HC1 and 
concentrations of copper and zinc were measured using flame atomic absorption 
spectrophotometry. Preparative and measurement techniques were calibrated against 
the National Bureau of Standards, Oyster Reference Material #1566. 



Apohemocyanin reconstitution experiments 

We have shown that the digestive gland of the American Lobster, Homarus ameri- 
canus, contains an abundant supply of copper-metallothionein (Engel and Brouwer, 
1986). The amino acid composition of the purified metallothionein from lobster is 
similar to that of the blue crab (Table I). In view of this similarity, and the relative 
ease with which it can be isolated from the lobster, we have used lobster digestive 
gland as the source of copper metallothionein in our apohemocyanin reconstitution 
experiments. 

Hemocyanin and copper metallothionein, to be used in copper transfer experi- 
ments, were prepared as described previously (Brouwer et al., 1986). Hemocyanin 
concentration was calculated from the optical density at 280 nm, using E}* m =: 14.3 
and a value of 75,000 for the molecular weight of a single oxygen-binding site carrying 
subunit (Nickerson and Van Holde, 197 1 ). Apohemocyanin was prepared by mixing 



242 D. W. ENGEL AND M. BROUWER 

hemocyanin in 50 mA/ Tris pH 8, 10 mMCaCl 2 , with an equal volume of buffer 
containing 20 rnM KCN. To prepare hemocyanin samples with different amounts of 
bound copper, the protein was either incubated with KCN for 10 minutes at room 
temperature, or dialyzed for 30 minutes against 20 mM KCN, followed by removal 
of the KCN on Sephadex G-25. Reconstitution of apohemocyanin was performed by 
mixing the apoprotein with purified copper metallothioneins in 50 mM Tris pH 8, 
10 mA/CaC^, in the absence of oxygen. 

Copper insertion into the active site of the apoprotein was measured by fluores- 
cence spectroscopy. Apohemocyanin was excited at 280 nm and the quenching of 
the tryptophan fluorescence, which accompanies copper incorporation, was moni- 
tored at 340 nm with a SPEX Fluorolog fluorescence spectrophotometer in the ratio 
mode. The concentration of functionally active oxygen binding sites was determined 
from the intensity of the copper-oxygen charge transfer band at 340 nm after addition 
of O 2 to the degassed incubation mixture. 

RESULTS 
Effect of overwintering on metal partitioning 

Differences were observed in the concentrations of copper in hemolymph and 
digestive gland samples among the three groups of intermolt C 4 crabs that were exam- 
ined (summer, 1985; winter-cold, 1986; and winter-warmed, 1986). In the hemo- 
lymph there was a correlation between the physiological condition of crabs and the 
concentrations of hemocyanin and copper (Fig. 1). Both summer and warmed hard 
crabs had hemocyanin and copper concentrations that were higher than the cold 
crabs, but only the difference between the copper concentrations in summer and cold 
hard crabs was significant (P < .05). Zinc concentrations did not change significantly 
(P > .05) among the three groups of crabs (Fig. 1 ), and did not appear to be positively 
correlated with hemocyanin concentration. In the digestive glands there was no sig- 
nificant difference (P > .05) in concentrations of copper between the summer and 
cold water crabs, but there was a significant (P < .05) increase in the crabs that were 
warmed (Fig. 2). Once again zinc concentrations did not show significant changes (P 
> .05). The large increase in copper concentration in the warmed crabs is correlated 
with the observed increase in hemocyanin in the hemolymph. 

The elution profiles obtained after gel-permeation chromatography of the cytosol 
from digestive glands of cold and warmed crabs showed differences in metals bound 
to metallothionein. Among the cold water crabs four of five had metallothionein 
peaks that contained primarily zinc, while three of four (i.e., one chromatographic 
sample lost) of the warmed crabs had metallothioneins that contained primarily cop- 
per (Fig. 3-II and III). Thus, the majority of cold water crabs had Cu/Zn ratios associ- 
ated with metallothionein that were reminiscent of premolt animals (i.e., high zinc 
low copper) while the majority of warmed crabs had patterns similar to those of sum- 
mer intermolt crabs (i.e., high copper low zinc) (Fig. 3-1). These data show that envi- 
ronmental conditions, physiological state, and feeding can affect tissue metal concen- 
trations and the cytosolic distributions of copper and zinc in blue crabs. 

Apohemocyanin reconstitution experiments 

Removal of copper from the active site of hemocyanin results in an increase of 
the intrinsic tryptophan fluorescence of the protein (Fig. 4). This observation allowed 
us to make a distinction between copper insertion and formation of native functional 



BLUE CRAB METAL METABOLISM 



243 



HEMOLYMPH 



o> 

E 



O 
O 

2 

LLJ 

I 



60 


. 


50 


- 


I 




1 


40 
30 


- 






I 




-- 




1 


20 


- 














10 


- 














n 

















o 

K 

QC ^ 



16.0 
14.0 



O o 10.0 

O E 8.0 
'o 

X 



o 



LU 
Q. 

a. 
O 
O 



6.0 
4.0 
2.0 



Z 
O 



LLI 

O 

Z 

N 



3.0 



o n 

2.0 



1.0 







I 



II III 



l-Summer Hard Crab 
ll=Winter Hard Crab (cold) 
Ill-Winter Hard Crab (warmed) 

FIGURE 1. Concentrations of hemocyanin, copper, and zinc in the hemolymph of blue crabs col- 
lected in the summer (Engel, 1987) and winter. The winter crabs all were collected at the same time. Half 
(5) were sampled at ambient temperature (6C) and the other half (5) were warmed to 20C and fed fish 
every other day for three weeks. Each histogram represents a mean of five individual crabs plus or minus 
standard error of the mean. 



oxygen binding sites. Both processes can be experimentally followed by fluorescence 
and absorbance spectroscopy as shown in Figure 5. The data demonstrated that incu- 
bation of apohemocyanin with copper metallothionein leads to fluorescence quench- 
ing before viable oxygen binding sites are formed, suggesting that copper insertion 
precedes the formation of biologically active oxygen binding sites (see Discussion). 

DISCUSSION 

As indicated earlier (Brouwer et ai, 1986), one of the proposed functions of cop- 
per metallothionein is as a Cu +1 donor for hemocyanin synthesis. The present experi- 
ment, with dormant and warmed crabs, shows that the predominance of copper on 
metallothionein among the warmed crabs is associated with the increased levels of 
hemocyanin in the hemolymph. Our earlier work with blue crabs also suggested a 
strong correlation between molting, copper metallothionein, and hemocyanin syn- 



244 



D. W. ENGEL AND M. BROUWER 



DIGESTIVE GLAND 



)NCENTRATiC 
mol/kg) 

^ en 0) -si 
b b b b 


- 




















b 3.0 


- 
















5 2.0 

fc 1.0 

8 


*- 


T 














_L 


1 




1 




II 


III 



7.0 
< *5 6.0 
fE^ 5.0 


- 


T 




I 




I 




1 


1 




I 


5 i 4.o 


- 














O T 
















ZQ 3.0 


- 














8x 2.0 


- 














o" 1.0 


- 














iTi n 

















l=Summer Hard Crab 
ll=Winter Hard Crab (cold) 
lll=Winter Hard Crab (warmed) 

FIGURE 2. Concentrations of copper and zinc in the digestive glands of blue crabs collected in the 
summer and winter. Further information on handling of the crabs is in Figure 1 . 



thesis (Engel, 1987). The studies reported in the present paper support that observa- 
tion and demonstrate that environmentally induced changes and nutrition also can 
cause changes in the copper/zinc ratios associated with metallothionein. This obser- 
vation is important because it further emphasizes the possible constitutive role of 
metallothioneins in normal metabolism. 

In the following section we will develop a model of the regulation of copper and 
zinc partitioning in the blue crab, based on studies by us (Engel, 1987; also present 
paper), Soumoffand Skinner ( 1 983), and Henry and Kormanik ( 1 985). 

The diagrams in Figure 6 display the physiological and biochemical processes 
involved in crustacean molting and the cyclic and chronological nature of these 
events. The first of these diagrams (Fig. 6 A) depicts the relative duration of the differ- 
ent portions of the molt cycle. The actual timing of events is dependent upon both 
environmental temperature and the size of the crab (Johnson, 1980). This type of 
presentation emphasizes the fact that the most dramatic/traumatic changes in the 
crabs occur over a relatively short period of time. The changes in concentrations of 
copper and zinc associated with metallothionein are dramatic and provide further 
evidence as to the dynamic nature of the molting process (Fig. 6B). If it is assumed 
that metallothionein-bound copper and zinc are associated with metalloprotein and 
metalloenzyme synthesis, we can predict when hemocyanin and zinc enzyme synthe- 
sis occurs during the cycle (Fig. 6C, D). These predictions are tentative and will need 
to be confirmed in future studies, since there are no direct data available in the litera- 
ture on these aspects of crustacean physiology. 

Copper metallothionein levels during the molt cycle are related directly to hemo- 
cyanin concentrations in hemolymph, and inversely to ecdysteroid concentrations in 
the hemolymph (Fig. 7). The decrease in digestive gland copper metallothionein is 



BLUE CRAB METAL METABOLISM 



245 



l-Summer Crab 



Q 

o 

111 
o 

z 
< 

CO 

<r 
O 

CO 

m 



Absorbance 
oCopper 




FRACTION NUMBER 



60 



O) 

3- 

o 

z 

N 
O 



cc 

LU 
CL 

CL 
O 
O 



FIGURE 3. Sephadex G-75 elution profiles of digestive gland cytosol prepared from blue crabs col- 
lected during the summer and winter. For further information on the crabs see Figure I. Protein separations 
were made using 60 mM Tris buffer, pH 7.9 with 2 mA/|tf-mercaptoethanol with a 2.6 X 60 cm column at 
a flow rate of 30 ml/h in all three groups of crabs (I, II, III). 



correlated with an increase in ecdysteroid liter in blue crab hemolymph (Soumoffand 
Skinner, 1 983). The ecdysteroid concentrations followed the same general pattern for 
males and immature females throughout the molt cycle. After molt the ecdysteroid 
level decreases rapidly with concomitant decreases in hemocyanin concentrations. 
Coincident with these decreases is an increase in copper metallothionein during the 
A 2 and B, stages, followed by an increase in hemocyanin during B, . Such interrela- 
tionships suggest a possible association between the molting hormone ecdysteroid, 
and the regulation of hemocyanin synthesis and levels of cytosolic copper. It is rele- 
vant to emphasize that synthesis of constitutive metallothioneins in mammals is un- 
der the control of steroid hormones (Karin et al, 1 980 a, b; Karin el ai, 1981). No 
such information exists for the invertebrate metallothioneins. The effect of the molt- 
ing hormone 20-hydroxyecdysone on metallothionein synthesis in the blue crab is 
presently under investigation. 

Comparisons of zinc metallothionein (Engel, 1987) and ecdysteroid concentra- 
tions (Soumoffand Skinner, 1 983) and carbonic anhydrase activity (Henry and Kor- 
manik, 1985) during the molt cycle suggest an inter-relationship between these three 



246 



D. W. ENGEL AND M. BROUWER 



9. 45X10- 



CO 

HI 



LU 

o 

LU 
O 

CO 

LU 

cc 
o 

13 



7.00X10- 




315.00 



335.00 
WAVELENGTH (nm) 



355.00 



FIGURE 4. Fluorescence intensity of deoxygenated lobster hemocyanin in 50 mM Tris pH 8.0 + 10 
mMCaC\ 2 as a function of the percentage copper remaining in the active site after dialysis against 20 mM 
Cyanide for 0, 5, 10, and 30 min. ( 1 ) 100%, (2) 60%, (3) 30%, (4) 8%. Excitation is at 280 nm. 



2.48 



CO 

z 

LU 



LU 
O 

z 

LU 
O 
CO 
LLJ 

rr 
O 



2.40 



2.32 



2.24 




.5 



3 







16 



24 



36 



TIME (hours) 

FIGURE 5. Change in fluorescence intensity and oxygen binding capacity (A Y) of partial apohemo- 
cyanin (5 nM) in 50 mM Tris pH 8.0 + 10 mM CaCl 2 still containing 35% of its original copper, as 
a function of incubation time with copper-metallothionein ( 10 ^M Cu) in the absence of oxygen. The 
fluorescence change (copper insertion) precedes the formation of viable oxygen binding sites. 



BLUE CRAB METAL METABOLISM 



247 



Hemocyanln 
Synthesis 

and 
Turnover 




Zinc Enzyme 
Synthesis 
(Carbonic 
Anhydrase) 




Activity Increasing 

FIGURE 6. Diagrammatic representation of the physiological and biochemical events occurring dur- 
ing the molt cycle of the blue crab. (A) the molt cycle of the blue crab with the duration of each portion 
indicating time. The designations of the molt stages are: C, * C 4 , hard crab; D, D 4 ; premolt; E, ecdysis; 
A,-A 2 , soft crab; B,-B 2 , papershell crab (Mangum, 1985). (B) The relative concentrations of copper and 
zinc on metallothionein are represented by the size of the copper, Cu and zinc, Zn symbols. (C) and (D) 
These two figures represent predicted hemocyanin and zinc enzyme synthesis activities generated from 
previously collected data (Engel, 1987). The degrees of shading are indications of the proposed activities 
of the biochemical pathways involved in hemocyanin synthesis and turnover, and zinc enzyme synthesis 
(carbonic anhydrase). 



components (Fig. 8). During the premolt period (D!-D 3 ) when both zinc metallothio- 
nein and ecdysteroid are at their peaks, the new epidermis is being synthesized be- 
neath the existing exoskeleton. At molt both zinc metallothionein and ecdysteroid 
decrease, and between stages A, and A 2 there is an abrupt increase in carbonic anhy- 
drase activity in the newly formed exoskeleton epidermis (Henry and Kormanik, 
1985). This rapid increase, which occurs over a period of hours, suggests that the 
enzyme may be synthesized and present in the new epidermis as an apo-protein, and 
is not activated by zinc until after molt. Even though the decrease in zinc-metallothio- 
nein occurs in the digestive gland and the increase of carbonic anhydrase in the epi- 
dermis, these two events may be linked. Possibly some of the zinc bound to metallo- 
thionein at the time of molt could be mobilized via the hemolymph to activate the 
apo-carbonic anhydrase. This hypothesis is attractive since preliminary results from 
our laboratory have shown the release of zinc from zinc metallothionein during stages 
A, and A 2 (D. W. Engel, unpub. data). 

The proposed cycles for copper and zinc-metallothionein (Figs. 7, 8) are specula- 
tive, but they are based upon the best available information on the physiological and 



248 



D. W. ENGEL AND M. BROUWER 



> z 
o o 



2 < 

at oc 



> o 

i- z 

< O 

- 1 o 

111 

or 



Ecdysteroid* 
Hemocyanin 
.Copper Metallothionein 




J I 



i L 



I I 



I I 



C D, D 2 D 3 D 4 E A, A 2 B, B 2 C 



MOLT STAGE 



ill 



x a: 

i- i- 

O z 

-I LU 



is 

til O 
Q. cr 

Q. LLJ 
O H 
O tO 



-I Q 

Z 
DC < 



*Soumoff and Skinner (1983) 



FIGURE 7. A diagrammatic representation of the processes involved in hemocyanin synthesis and 
turnover, and the interactions between ecdysteroids and copper metallothionein. The data on hemocyanin 
and copper metallothionein concentrations are from Engel ( 1987) and for ecdysteroid from SoumorTand 
Skinner (1983). 

biochemical events controlling metal partitioning during molt. These changes are 
reproducible, and our experiments concerning the effects of thermal changes on 
metal distributions give further support to the hypothesis that metallothionein is a 
constitutive metal-binding protein in blue crabs. 



LLJ 

to 
oc 

Q 



CD h- 

cr o 
o 

LLJ 



LLJ 

Lt 



Carbonic Anhydrase* 

Zinc Metallothionein 

Ecdysteroid** 




""r" V 



D 2 D 3 D 4 A, 
MOLT STAGE 



A 2 



Bi 






LLJO 



ILJ 



LU 
DC 



tULLJ 



LLJO 
DC HI 



B 2 



*Henry and Kormanik (1985) **Soumoff and Skinner (1983) 

FIGURE 8. A diagrammatic representation of the processes involved in the synthesis of zinc-depen- 
dent enzymes and in particular carbonic anhydrase, and how zinc metallothionein and ecdysteroid interact 
to affect enzyme activity. The data on zinc metallothionein are from Engel (1987), on ecdysteroid from 
SoumorTand Skinner (1983). and carbonic anhydrase from Henry and Kormanik (1985). 



BLUE CRAB METAL METABOLISM 249 

The events and processes described here and in an earlier publication (Engel, 
1987) do not address the question of control of the molt cycle and metal turnover. 
During the molt cycle there are pronounced changes in the cytosolic distribution 
and tissue concentrations of metals and accompanying changes in the hemolymph 
ecdysteroid concentrations. Studies by Singer and Lee (1977) suggest that the hemo- 
lymph hormonal levels are modulated by changing MFO (mixed function oxygen- 
ases) activity in the antennal gland. These authors demonstrated that MFO activity 
in the gland also varies with stages of the molt cycle. This activity is negatively corre- 
lated with the ecdysteroid levels (Soumoffand Skinner, 1983), suggesting that the 
MFO system controls steroid/hormonal concentrations in molting blue crabs, which 
in turn may affect metal partitioning as described in this paper. 

Further evidence for metallothionein's metal regulatory function comes from the 
in vitro hemocyanin reconstitution experiments. Apohemocyanin can only be recon- 
stituted with Cu +1 (Konings et a/., 1969; Lontie and Witters, 1973). Since copper 
binds to metallothionein as Cu +l , and since copper metallothionein levels and hemo- 
cyanin biosynthesis seem to be linked in vivo, we initiated a study of hemocyanin- 
copper metallothionein interaction in vitro. The data presented in Figure 4 show that 
the intrinsic tryptophan fluorescence of lobster hemocyanin strongly depends on the 
amount of Cu +1 bound to the active site. This is in line with the observations that 
several crustacean hemocyanins contain tryptophan residues in close proximity to 
the binuclear copper site (Gaykema et al., 1984; Linzen et ai, 1985). This property 
allowed us to make a distinction betwen Cu +1 incorporation into the active site of 
apohemocyanin and the formation of native functional oxygen binding sites. It is 
evident from Figure 5 that the quenching of tryptophan fluorescence, observed when 
apohemocyanin is incubated with copper metallothionein, precedes the formation of 
biologically active oxygen binding sites. This strongly suggests that the copper transfer 
process is followed by a slow reordering of the tertiary structure of the copper sites to 
the native configuration. Similar sequences have been demonstrated for the reconsti- 
tution process of many Cu +2 proteins where binding of copper to the active site is 
followed by a slow return of the protein to its biologically active state (Kertesz et al., 
1972; Morpurgo et al., 1972; Rigo et al., 1978; Marks et al., 1979; Blaszak et al., 
1 983). This observation may also explain why reconstitution of apohemocyanin with 
copper metallothionein can only be accomplished under anaerobic conditions 
(Brouwer et al., 1986). The Cu +1 in the distorted sites is not capable of combining 
reversibly with oxygen. Interaction of oxygen with Cu 41 under these conditions re- 
sults in oxidation of metal. These Cu +2 -sites will not bind oxygen and are lost for 
detection by absorbance spectroscopy. This hypothesis is presently under further in- 
vestigation. 

The studies described in this paper have demonstrated that marine Crustacea are 
excellent model systems to study the role of metallothionein in copper/zinc metabo- 
lism on an organismal, cellular, and molecular level. Only when this function of met- 
allothionein is fully understood will it be possible to assess its value as a metal-detoxi- 
fying protein. 

ACKNOWLEDGMENTS 

The authors thank Mr. William J. Bowen III, and Lt. (jg) Debra Davis of our 
Laboratory for their assistance during this investigation. Hooper Family Seafood, 
Smyrna, NC; and Pittman Seafood, Merrimon, NC, supplied the crabs used in this 
investigation. The authors also thanks Dr. Bruce A. Fowler, National Institute of 



250 D. W. ENGEL AND M. BROUWER 

Environmental Health Sciences; Dr. G. Roesijadi, Chesapeake Biological Labora- 
tory, University of Maryland; and Drs. Brenda Sanders and Kenneth Jenkins, Cali- 
fornia State University, Long Beach, for reviewing this manuscript. 

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COUSINS, R. J. 1982. Relationship of metallothionein synthesis and degradation to intracellular zinc me- 
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GEORGE, S. G., AND A. VIARENGO. 1985. A model of heavy metal homeostasis and detoxification in 
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HENRY, R. P., AND G. A. KORMANIK. 1985. Carbonic anhydrase activity and calcium deposition during 
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JOHNSON, P. T. 1 980. Histology of the Blue Crab, Callinectes sapidus. A Model for Decapoda. Praeger Sci., 
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KARIN, M., H. R. HERSCHMAN, AND D. WEINSTEIN. 1980a. Primary induction of metallothionein by 
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KARIN, M., R. D. ANDERSEN, E. SLATER, K. SMITH, AND H. R. HERSCHMAN. 1980b. Metallothionein 
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phys. Acta 194: 55-66. 

LINZEN, B., N. M. SOETER, A. F. RIGGS, H. J. SCHNEIDER, W. SCHARTAU, M. D. MOORE, E. YOKOTA, 
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LONTIE, R., AND R. WITTERS. 1973. Hemocyanin. Pp. 334-358 in Inorganic Biochemistry, Vol. I, G. L. 
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BLUE CRAB METAL METABOLISM 25 1 

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Reference: Biol. Bull. 173: 252-259. (August, 1987) 



FREE D-AMINO ACIDS IN THE TISSUES OF MARINE BIVALVES 

HORST FELBECK AND SANDRA WILEY 

University of California San Diego, Scripps Institution oj Oceanography, Marine Biology 
Research Division, A-002, La Jo/la, California 92093 

ABSTRACT 

Seventeen species of marine bivalves were surveyed for the presence of free D- 
alanine, D-aspartate, and D-valine in their tissues. D-aspartate was found in several 
species in concentrations approaching those of L-aspartate. D-alanine was detected 
particularly in lucinid and vesicomyid clams at levels exceeding manyfold those of 
L-alanine. D-valine was absent in all cases. A test of a hydrolysate of bulk soluble 
proteins of Lucinoma aequizonata, a species characterized by extremely high levels 
of D-alanine, showed no major incorporation of D-alanine into proteins. The im- 
plications of these results, for previously published analytical data and for human 
nutrition, are discussed. 

INTRODUCTION 

D-amino acids generally are viewed as natural oddities. They are usually not 
found in proteins, and they occur only occasionally in sizable quantities, either freely 
dissolved, or incorporated into peptides and metabolites in the tissues of animals and 
plants (Robinson, 1976; Bodanszky and Perlman, 1969; Corrigan, 1969). However, 
it has been suggested that the D-isomers of amino acids were as common as the L- 
forms in ancient prebiotic times on earth. Since the L-isomers were used exclusively 
by various life forms, they were removed from this natural equilibrium. The supply 
of L-amino acids was maintained by chemical racemization from the D-isomers left 
behind (Aono and Yuasa, 1977). Even today relatively large amounts of D-amino 
acids can be identified in oceanic waters, where apparently they have been formed 
by chemical racemization from the large pool of dissolved L-amino acids (Lee and 
Bada, 1977). 

Widespread attention was first focused on D-amino acids when they were pro- 
posed to be causes and indicators of cancer. Proteins in tumor tissues were thought 
to contain high levels of D-glutamic acid, thus distinguishing them from normal tis- 
sue (Koegl and Erxleben, 1939). This theory was rejected, however, after years of 
controversy (Miller, 1950). 

Recently, the occurrence of D-aspartate instead of the L-form in some proteins 
has been a focus of investigation for molecular repair mechanisms (McFadden and 
Clarke, 1982). It was reported that methylated aspartyl residues in erythrocyte mem- 
brane proteins had been converted to the D-form. According to the theory, the ap- 
pearance of D-aspartate in proteins is a first sign of degradation. These proteins are 
either tagged for disposal, or the D-aspartyl residue can be reversed to the L-form 
after methylation. 

A similar mechanism, time-dependent chemical racemization of aspartate within 



Received 9 September 1986; accepted 28 May 1987. 

252 



D-AMINO ACIDS IN BIVALVES 253 

proteins with a low turnover rate, like eye lens or dental proteins, has been used to 
date these proteins by measuring the ratio of the D- to the L-form of aspartate (Mas- 
ters, 1983; Bada and Brown, 1980). Exactly the same principle of racemization has 
also been used extensively to determine, with great accuracy, the ages of fossils, since 
the normally present L-aspartate racemizes chemically at a constant rate after the 
death of an organism (Bada and Schroeder, 1975). 

D-aspartate may also be a neurotransmitter (Wiklund et ai, 1982). Presently, it 
is being used experimentally as a non-metabolizabie replacement for L-glutamate 
and L-aspartate in neurotransmitter research, since it can use the same uptake sites 
(Drejer<Y0/.. 1983; Taxt and Storm-Mathisen, 1984). 

Since finding that the artificial sweetener aspartame produces D-aspartate when 
heated (e.g., during cooking) another line of research has been initiated (Boehm and 
Bada, 1 984). Thus humans may be exposed nutritionally to considerable amounts of 
a D-amino acid due to increased consumption of aspartame worldwide. 

Only a few publications have focused on the metabolic role of free D-amino acids 
in animal and plant tissue. D-alanine is present in some molluscs (Matsushima et a/., 
1984) and crustaceans (D'Aniello and Giuditta, 1980), and it has been demonstrated 
to be synthesized during anaerobic metabolism in annelids (Felbeck, 1 980; Schoettler 
et ai, 1983). Recently, a study of D-amino acids, as indicated by the reaction with 
D-amino acid oxidase, in a variety of marine invertebrates was published (Preston, 
1987a). D-amino acids were found in 18 of the 43 species of the 8 phyla surveyed. 
The presence and metabolism of D-aspartate has been investigated in the tissues of 
the bivalve Solemya reidi (Felbeck, 1985) and of some cephalopods (D'Aniello and 
Giuditta, 1 977, 1 978). In all cases, the D- and the L-forms were present in about equal 
concentrations. The concentrations of D- and L-alanine in the polychaete Arenicola 
marina are approximately the same, and the concentrations of the two isomers in- 
crease similarly in response to metabolic stress (Felbeck, 1 980; Schoettler et ai, 1 983). 
The bivalve Solemya reidi takes up D-aspartate from environmental seawater and 
metabolizes it just as quickly as it does the L-form. Initially the D-form is converted 
into the L-isomer before further metabolism takes place (Felbeck, 1985). The uptake 
and metabolism of D-alanine from seawater has been described recently for coelomo- 
cytes of the annelid Glycera dibranchiata (Preston, 1987b). 

Marine invertebrates commonly have extremely high concentrations of free 
amino acids which, in the event of osmotic stress, form the largest share of the pool 
of intracellular osmolytes (Bishop et ai, 1983). Therefore, D-amino acids in this pool 
might serve as important metabolic reserves, sinks, or regulatory factors. 

Several investigators have described either isolated occurrences of individual D- 
amino acids in some invertebrates or have measured the unspecified presence of D- 
amino acids. No known publication has surveyed organisms for individual D-amino 
acids. The recent availability of chromatographic screening techniques for some 
amino acid isomers prompted our investigation of a number of marine bivalves for 
the presence of specific D-amino acids. We chose to study the Bivalvia because their 
physiology is well known, a variety of species is readily available, and their tissues 
contain high concentrations of free amino acids. 

MATERIALS AND METHODS 
Animals 

Animals were purchased live at fish markets or collected from a variety of loca- 
tions (Table I). The animals from the Santa Barbara channel were collected shipboard 



254 



H. FELBECK AND S. WILEY 



TABLE I 



Collection areas of animals 



Animal 



Collection area 



Bathymodiolus thennophilus 
Calyptogena elongata 
Cliione californiensis 
Chione stiitchburyi 
Codakia obicularis 
Codakia tigerina 
Corbicula (liiminea 
Crassostrea virginica 
Hiatel/a pholadis 
Hi unites multirugosus 
Lima hemphilli 
Lucinoma aequizonata 
Mercenaria mercenaria 
Modiohis capax 
Mytilus edulis 
Solemya reidi 
Tapes japonic a 



Pacific, Galapagos hydrothermal vents 

Pacific, Santa Barbara channel 

Pacific, Gulf of California 

Fish market, San Diego, California 

Atlantic, Bahamas, intertidal 

Fish market, Phillipines 

Fish market, San Diego 

Fish market, San Diego 

Pacific, La Jolla, California 

Pacific, San Diego 

Pacific, San Diego 

Pacific, Santa Barbara channel 

Fish market, San Diego 

Pacific, San Diego 

Fish market, San Diego 

Pacific, Santa Monica Bay 

Fish market, San Diego 



by otter-trawl. Solemya reidi was collected shipboard in Santa Monica Bay by Van 
Veen grab. Most other species were collected by divers and were maintained alive in 
flow-through seawater tanks, at approximate in situ temperatures, for a maximum of 
ten days before being sacrificed. Bathymodiolus thermophilus specimens were col- 
lected by the submarine DSRV "Alvin" during a cruise to the Galapagos hydrother- 
mal vents. The animals were frozen upon retrieval. Codakia tigerina was purchased 
alive at a fish market in the Philippines and then shipped by air freight in 70% alcohol. 

Sample preparation 

To account for the presence of symbiotic bacteria in the gills of some of the bi- 
valves used in this study (Felbeck el a/., 1981; Felbeck, 1983), all bivalves were 
opened, and the gills were removed and analyzed separately from the remaining 
soft parts. 

The tissue, frozen with liquid nitrogen, was first pulverized in a mortar. The ho- 
mogenization was then completed in 1 N HC1O 4 with an Ultra-Turrax homogenizer. 
The homogenate was centrifuged at 12,000 X g for 15 min, and the supernatant was 
neutralized with 3 M KHCO 3 . The resulting precipitate was removed by centrifuga- 
tion. An aliquot of this extract was derivatized with o-phthaldialdehyde (OPA) and 
N-acetyl-L-cystein (NAC), according to the method described by Aswad (1984). The 
amino acid isomers were then separated on a CIS reverse phase column with a gradi- 
ent of 50 mA/ sodium acetate, pH 5.8, containing 8% methanol (Sol. A) to methanol 
(Sol. B). The gradient was (in % of solution B): min, 0%; 4 min, 0%; 10 min 25%; 
20 min 27%; 34 min, 52%; and 50 min, 52%. Using this gradient which was modi- 
fied from the one described by Aswad (whose sole purpose was to separate D-and L- 
aspartate) the two alanine and valine isomers could also be separated completely. 

Using a Gilson Datamaster integrator, standards for the D- and L-isomers of 
aspartate, valine, and alanine were used to determine standard response curves. 
When samples were analyzed, the area under each individual peak was used to deter- 
mine concentration and, subsequently, the ratio of the individual stereoisomers. 



D-AMINO ACIDS IN BIVALVES 255 

To determine whether D-alanine was present in the proteins of L. aequi-onata, 
tissue of a whole animal was homogenized in distilled water with an Ultra Turrax. 
After centrifugation, the pellet was twice resuspended and rehomogenized in water. 
The combined supernatants were dialyzed against multiple changes of distilled water 
for five days to remove all free amino acids. The resulting solution of mixed soluble 
proteins of L. aequizonala was then precipitated with perchloric acid, centrifuged, 
and the pellet hydrolyzed overnight with HC1. The hydrolyzate was then analyzed 
for D-amino acids as described above. 

RESULTS 

Significant concentrations of D-aspartate and D-alanine and their L-isomers were 
detected (Table II); no D-valine was found. All Lucinidae showed high concentra- 
tions of D-alanine concentrations much higher than those of L-alanine. D-alanine 
also was detected in Mercenaria mercenaria, both species ofChione, Hinnites gigan- 
teus, Lima hemphilli (gills), Bathymodiolus thermophilus, Crassostrea virginica 
(gills). Tapes japonica, Hiatella pholadis, and Corbiculafluminea. In all of these spe- 
cies, the concentration ratios of D- to L-isomer was below one. In the Mytilidae Myti- 
lus edluis and Modiolus capax, no D-alanine was detected, but D-aspartate was found 
in concentrations approaching those of the L-isomer. D-aspartate was also detected 
in the gills of Bathymodiolus thermophilus. 

No D-alanine was detected in the hydrolyzed soluble protein fraction of Luci- 
noma aequi~onata. 

Because amino acid levels among individual animals of the same species are typi- 
cally highly variable, we did not attempt to establish average concentrations for a 
large number of bivalves but instead focused on the presence of D-amino acids. We 
postulate that the detection of D-amino acids in any individual organism is significant 
for the species in general. 

DISCUSSION 

The lucinids contained the highest D- to L-ratio of alanine. The extremely high 
level of free alanine in Codakia ohicularis tissues has been measured only by ion- 
exchange amino acid analysis and, therefore, has been attributed entirely to "generic" 
alanine acting as an osmoregulatory agent or an end-product of anaerobic metabo- 
lism (Berg and Alatalo, 1984). In fact, most of this alanine is in the D-form, prompt- 
ing us to question the function of the D-alanine in this bivalve as well as in all lucinid 
clams. One possibility is that the D-alanine is entirely made by the symbiotic bacteria 
inside the cells of the gill. Gram negative bacteria, like the symbiotic species found in 
the gill (see Schweimanns and Felbeck, 1 985, for review), often contain D-alanine in 
their cell walls (Katz and Detrain, 1977). Therefore, extraction of the cell wall could 
yield significant amounts of the D-isomer of alanine. In addition, these bacteria are 
thought to provide a major share of the bivalves' nutritional needs by fixing CCK from 
the seawater and transferring reduced organic compounds, possibly including D-ala- 
nine, to the host. It is unlikely that the D-amino acids originate in bacteria, however, 
since tissues lacking bacteria have a D- to L-isomer ratio similar to that of gills densely 
populated with bacteria. If the bacteria produce and export D-alanine, then the gill 
preparations should show a larger share of the D-isomer. In addition, some bivalve 
species (Table II) without symbiotic bacteria also have high concentrations of D- 
alanine. 

Another peculiar aspect of the large proportion of D-alanine in the free amino 



256 



H. FELBECK AND S. WILEY 



TABLE II 

Concentrations ofD- and L-amino acids in the tissues of marine bivalves 



Animal 


Tissue 


n 


L-alanine 
(umol/g 
fw) 
(xSD) 


D-alanine 
(umol/g 
fw) 
(x + SD) 


Ratio 
D/L 


L-aspartate 
(umol/g 
fw) 
(x + SD) 


D-aspartate 

( umol/g fw) 
(xSD) 


Ratio 
D/L 


SOLEMYIDAE 


















Solemya reidi 


foot 


2 


14.2 6.9 


0.1 0.1 


0.01 


13. 3 2.4 


12.4 1.6 


0.93 




gill 


2 


3.9 0.7 


1.4 0.6 


0.36 


7.6 2.0 


5.4 2.2 


0.71 


MYTILIDAE 


















Mytilus edulis 


foot 


2 


4.4 + 0.2 








4.6 1.8 


1.4 0.5 


0.3 




gill 


2 


2.8 0.9 








3.9 0.8 


3.5 0.7 


0.9 


Modiolus capax 


foot 


2 


2.7 1.4 


4 5.7 


1.5 


6.9 + 2.7 


2.6 0.01 


0.38 




gill 


1 


0.5 


0.7 


1.4 


2.9 


2.3 


0.79 


Bathymodiolus 


foot 


1 


12.3 


0.9 


0.08 


2.4 








thermophilus 


gill 


1 


11.4 


3.2 


0.28 


2.2 


0.7 


0.31 


OSTREIDAE 


















Crassostrea virginica 


mantle 


1 


19.4 








3.5 










gill 


2 


9.9 2.6 


1.1 0.1 


0.12 


5. 3 0.5 








PECTINIDAE 


















Hinnites 


foot 


2 


1.4+ 1.2 


1.1+0.9 


0.79 


1.0 1.3 


0.4 0.6 


0.4 


multirugosus 


gill 


2 


0.5 0.2 


0.7 0.9 


1.4 


0.5 0.2 


0.2 0.2 


0.4 



LIMIDAE 

Lima hemphilli 
VENERIDAE 



gill 



1.0 + 0.5 



0.5 0.3 



0.5 



0.8 0.1 



Tapes japonica 


foot 


1 


13.4 


11.2 


0.84 


15.5 










gill 


1 


4.7 


3.5 


0.74 


1.7 


0.2 


0.12 


Chione califomiensis 


foot 


1 


3.0 


2.1 


0.7 


14.6 










gill 


3 


3.8 2.6 


1.8+ 1.5 


0.47 


4.5 0.9 








Chione stutchburyi 


foot 


1 


4.6 


2.0 


0.44 


6.6 










gill 


2 


3.3 1.5 


1.3 0.7 


0.39 


3. 3 2.8 








Mercenaria 


foot 


2 


17.7 + 3.1 


16.2 1.9 


0.92 


7.8 3.5 








mercenaria 


gill 


3 


4.2 2.2 


2.8 1.4 


0.67 


4.1 + 1.4 








CORBICULIDAE 


















Corhiaila /Iiiminea 


foot 


2 


1.9 0.4 


0.9 1.2 


0.47 


0.7 0.2 










gill 


3 


3.8 0.8 


0.6 + 0.1 


0.16 


0.4 0.3 








HIATELLIDAE 


















Hiatella pholadis 


gill 


2 


4.4+ 1.0 


1.4 0.2 


0.32 


2.3 .01 








VESICOMYIDAE 


















Calyptogena 


foot 


1 


1.1 


11.3 


10.45 


11.5 








elongata 


gill 


1 


0.3 


2.7 


9.0 


1.4 








LUCINIDAE 


















Codakia obicularis 


foot 


1 


13.5 


187.2 


13.9 


0.3 


0.2 


0.67 




gill 


1 


2.4 


26.6 


li.l 


0.3 


0.1 


0.33 


Codakia ligerina 


foot 


1 


0.4 


22.4 


56.0 


0.6 


0.1 


0.17 




gill 


1 


0.4 


21.5 


53.7 


0.4 








Lucinoma 


foot 


1 


3.8 


84.2 


22.2 


0.4 


0.3 


0.8 


aequizonala 


gill 


3 


3.1 0.5 


26.1 4.1 


8.4 


1.0 0.9 


0.1 


0.1 



The levels are given in ^mol/gram fresh weight with the standard deviation. When only gills were tested, the foot was 
too small to be easily dissected and analysed. (" " below 0. 1 ^mol/g fw) 



acids of Lucinidae is that the enzyme most commonly responsible for the formation 
of D-amino acids amino-acid racemase would cause an equal distribution be- 
tween the two isomers (Barman, 1969). The fact that up to 98% of the free alanine 
pool is in the D-form suggests that: (a) another specialized enzyme is responsible 
for the metabolism of the D-isomer; and (b) the D-isomer is not metabolized after 
conversion to the L-form by a racemase, but used separately. Aside from the lucinid 
clams, however, other examples were found where the ratio of D- to L-alanine was 
lower: below one. Here we assume that a racemase interconverts the two isomers. 
The occurrence of D-aspartate can be explained more easily by the presence of a 



D-AMINO ACIDS IN BIVALVES 257 

racemase, for the maximal ratio of the D- to the L-isomer was around one. Indeed, 
this specific racemase has already been demonstrated in Solemya reidi (Felbeck, 
1 985). The aspartate-racemase does not catalyze the conversion of alanine. 

In spite of the high D-alanine concentration in Lucinoma aequizonata, no detect- 
able quantities of D-alanine were found in the proteins of this animal. Therefore, the 
selection for the L-isomer of alanine in protein synthesis is significant. Since the 
method used only provides a crude overview of a selected group of proteins those 
soluble in distilled water we cannot exclude the possibility that some minor fraction 
of the soluble or insoluble proteins would include D-alanine; neither of these would 
have been detected by the method used. 

Wide ranging surveys for the presence of D-amino acids are rarely in the literature. 
The review article by Corrigan (1969) and Preston's (1987a) recent results are the 
only known examples. Certainly, one reason is that simple, quick methods to deter- 
mine the concentrations of D- and L-isomers of individual amino acids have been 
published only recently. Before this, either both isomers were detected as a sum (e.g., 
in HPLC with OPA/mercaptoethanol derivatization or with the classical ion ex- 
change amino acid analyzer) or just the L-isomer was detected in typically stereospe- 
cific enzymatic determinations. Since it was always assumed that no D-amino acids 
were present, the results obtained by these methods were taken as representative for 
"all" amino acids. The D-amino acids concentrations found by Preston ( 1 987a) were 
obtained unspecifically with a test using D-amino acid oxidase and, therefore, were 
only applicable as indicator of the general presence of most D-amino acids ( D-aspar- 
tate and D-glutamate do not react with the D-amino acid oxidase). 

Our survey includes the amino acids alanine and aspartate, both of which are 
commonly found in high concentrations in marine invertebrates, and shows the fre- 
quent occurrence of both stereoisomers. 

This result is significant for "standard" experimental research organisms like Myt- 
ilus edulis (Bishop et ai, 1 983). In this species, the pool of free aspartate is used as an 
initial substrate for anaerobic metabolism (see de Zwaan and Putzer, 1985, for re- 
view). Whenever the concentration of this amino acid was tested using enzymatic 
methods, only about half of the available amino acid was detected; i.e., the pool of 
aspartate was actually higher than measured. This may explain the apparent lack of 
enough initial substraate for anaerobic energy metabolism, as recently reviewed by 
de Zwaan and Putzer (1985). Similarly, in other organisms such as the polychaete 
Arenicola marina, the initial depletion of the (enzymatically measured) L-aspartate 
is not large enough (Felbeck, 1980; Schoettler et ai, 1983). We think it is possible 
that D-aspartate, as well as the L-isomer, occurs in Arenicola, and that it serves there 
as additional substrate not detected by enzymatic analysis which after rapid racemiza- 
tion also can be used as metabolic substrate. 

We conclude that many published results where amino acid levels in invertebrates 
have been used as indicators for metabolic pathways, or to calculate metabolic rates, 
will have to be reassessed because D-amino acids may be present in the tissues used. 

Currently, we can only speculate what the metabolic role of D-amino acids is in 
marine invertebrates. Amino acids are usually used as osmolytes in the tissues of 
marine invertebrates and, therefore, are often present in very high concentrations 
(Bishop, 1983; Yancey et al, 1983). The exchange of part or most of the L-isomer 
for the D-isomer may influence regulatory mechanisms involving these amino acids. 
Glutamate-pyruvate-transaminase is inhibited by high levels of L-alanine (Barman, 
1969); the D-isomer may not have this effect on this enzyme. 

Finally, large quantities of free D-amino acids in tissues of common marine bi- 



258 H. FELBECK AND S. WILEY 

valves may affect human health. Some of our test species were obtained from com- 
mercial fish markets. For example, large quantities of D-alanine were found in Co- 
dakia tigerina specimens bought in a fish market in the Philippines. These fish are 
routinely consumed by humans. Even Mytilus edulis, one of the most common bi- 
valves cultured and consumed in large quantities, contains a concentration of D- 
aspartate, equal to that of L-aspartate, which is usually between 3 and 14 ^mol/g 
fresh weight (de Zwaan and Putzer, 1985). Little research has been done on the me- 
tabolism of D-amino acids in humans or on the effect of long-term exposure to D- 
amino acids. D-amino acids can cause analgesia in humans, and some D-amino acids 
are powerful inhibitors of some enzymes involved in regular metabolic pathways 
(Koyuncuoglu and Berkman, 1982). The result presented in this paper that some 
D-amino acids exist sometimes in extremely high concentrations in commonly con- 
sumed shellfish should prompt a closer examination of the effects of D-amino acids 
on humans. 

ACKNOWLEDGMENTS 

This research was funded by NSF grant OCE83- 1 1 259 to HF and George Somero. 
We thank Ron McConnaughey and John O'Sullivan from SIO, and Mr. Colin Higgs 
of the Department of Fisheries (Nassau, Bahamas) for collecting and providing speci- 
mens. Spencer Luke for identifying most of the species, and Drs. Patricia Masters 
and JeffBada for help in hydrolyzing a protein sample and assistance with the HPLC. 
Mr. Robert Yin kindly provided Codakia tigerina from the Philippines. 

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Reference: Biol. Bull. 173: 260-276. (August, 1987) 



TROPHOSOME ULTRASTRUCTURE AND THE CHARACTERIZATION 
OF ISOLATED BACTERIOCYTES FROM INVERTEBRATE-SULFUR 

BACTERIA SYMBIOSES 

STEVEN C. HAND 

Department of Environmental, Population and Organismic Biology, University of Colorado, Campus Box 

B-334. Boulder, Colorado 80309. and Department of Biology. University of 

Southwestern Louisiana, Lafayette, Louisiana 70504 

ABSTRACT 

Electron microscopy of trophosome tissue from the vestimentiferan tubeworm 
Riftia pachyptila clearly indicates that the bacterial symbionts are enclosed within 
animal cells (bacteriocytes). The structure of this lobular tissue is complex. Each lob- 
ule consists of an outer layer of trophochrome cells (devoid of symbionts, but with 
numerous pigmented granules), an inner region of bacteriocytes, and a central hemo- 
lymph space. Sulfur deposits within bacteria decrease in size and number with in- 
creasing distance of the bacteria from the hemolymph space. Bacteria located toward 
the center of the lobule appear smaller than those nearer the periphery, suggesting 
that metabolic and developmental gradients exist. Trophochrome cells and free bac- 
teria were enriched from the trophosome ofR. pachyptila. 

A procedure is described for the isolation of bacteriocytes from gill tissue of the 
bivalves Calyptogena magnified and Lucina floridana. Numerous bacteria reside in 
vacuoles within the bacteriocyte cytoplasm, as do large (5- 10 micron), heterogeneous 
granules. Maximum CO 2 fixation rate at 20C for bacteriocytes from C. magnified is 
13.2 nmoles CO 2 /mg protein/h, compared to 21.6 nmoles CO 2 /mg protein/h for L. 
floridana bacteriocytes. Fixation by bacteriocytes from C. magnified is inhibited by 
sulfide, and to a lesser extent thiosulfate, at 0.1-1.0 mA/. Thiosulfate increases CO 2 
fixation two-fold in L. floridana bacteriocytes. 

C. magnified bacteriocytes incubated for 1 h in 0.5 mA/ sulfide maintain higher 
intracellular ATP concentrations (3.3 nmoles/million cells; 1.01 mA/) than do con- 
trol cells without sulfide (1.02 nmoles/million cells; 0.31 mA/). These results and 
comparable observations suggest that the identities of exogenous sulfur compounds 
exploited for chemical energy by the symbiosis may depend on the structural integrity 
and organization of the experimental preparation. 

INTRODUCTION 

In symbiosis between sulfur bacteria and marine invertebrates, various metabolic 
features are critically dependent on the cellular integrity of each participant. To study 
these characteristics without disrupting cellular structure, we developed a procedure 
for isolating intact bacteriocytes (eucaryotic cells that contain large numbers of bacte- 
rial endosymbionts) from gill tissues of the hydrothermal vent clam, Calyptogena 
magnified, and the shallow-water bivalve Lucina floridana, an inhabitant of seagrass 
beds. An ultrastructural description of the intact trophosome of the hydrothermal 

Received 16 April 1987; accepted 29 May 1987. 

260 



INVERTEBRATE-BACTERIA SYMBIOSES 261 

vent tubeworm Riftiapachyptila (Pogonophora) provides new information about cel- 
lular arrangements and metabolic potentials in this symbiont-containing tissue. Fi- 
nally, using bacteriocyte suspensions prepared from the bivalves, we measured intra- 
cellular ATP levels and the capacity for carbon fixation in the presence of various 
sulfur compounds. 

The primary advantage of using bacteriocytes for metabolic studies is that the 
symbiotic bacteria are retained in their natural microenvironment. As a conse- 
quence, the bacteria receive chemical signals (sulfur compounds, dissolved gases, etc.) 
via the cytoplasm of the host cell. Furthermore, all bacteriocyte surfaces are in direct 
contact with medium constituents, so that the effects of slowly exchanging compart- 
ments such as connective tissue spaces are minimized. Thus individual bacteriocytes 
are considered functional symbiotic units, and their response to various stimuli quan- 
tified on a cellular basis. 

Until now, isolated invertebrate cells have not been used to study physiological 
and biochemical relationships between sulfur oxidizing bacteria and the host. Rather, 
previous studies have focused on other levels of biological organization and complex- 
ity. Data have been obtained using ( 1 ) the intact symbiosis at the whole-organism 
level (e.g., Anderson, 1986; Arp et aL 1984; Childress et al., 1984; Felbeck, 1983, 
1985; Fiala-Medioni et al.. 1986), (2) excised tissues (e.g., Cavanaugh, 1983; Dando 
etal, 1985; Felbeck, 1983; Powell and Somero, 1983), (3) variously prepared homog- 
enates of tissues (e.g., Felbeck, 1981; Felbeck etal., 1981; Fisher and Childress, 1984; 
Fisher and Hand, 1984; Hand and Somero, 1983; Powell and Somero, 1985, 1986a), 
and (4) isolated bacteria and cellular organelles (e.g., Belkin et al., 1986; Powell and 
Somero, 1986b). Depending on the degree of tissue disruption, significant variation 
was observed in the metabolism of sulfur compounds and the rates and characteristics 
of carbon fixation. For example, using homogenates of gill tissue from C. magnifica, 
Powell and Somero (1986b) reported that stimulation of ATP synthesis by sulfur 
compounds occurred only when bacteria contained therein were lysed. 

Experimental preparations that maintain the bacteria in more biologically realis- 
tic surroundings offer new opportunities for assessing their metabolic potential. This 
possibility was the impetus for the present study. In addition to isolating bacteriocytes 
from gill tissue of C. magnifica and L. floridana, we isolated trophochrome cells 
(green pigmented cells) and free bacteria from trophosome tissue of R. pachyptila. 
Although electron micrographs presented herein indicate bacteriocytes within the 
trophosome, we were unsuccessful in isolating them intact from this source. 

MATERIALS AND METHODS 
Experimental animals and reagents 

Specimens of Rift i a pachyptila and Calyptogena magnifica were collected in 
March 1985 during the hydrothermal vent expedition to the Galapagos Rift with the 
submersible DSRV Alvin at the "Rose Garden" 1 site (Hessler and Smithey, 1983). 
The live animals on board the RV Melville were handled as described by Powell and 
Somero ( 1986a). Tissue samples were dissected from specimens ofR. pachyptila and 
C. magnifica typically within 3 h of their arrival on board ship. Tissue weights were 
determined with the motion compensated shipboard balance developed by Childress 
and Mickel( 1980). 

Specimens of the eulamellibranch bivalve Lucina floridana were collected from 
the sulfide-rich sediments of Thalassia and Ruppia seagrass beds in St. Joseph's Bay, 
Florida. Animals were maintained in the laboratory as described by Fisher and Hand 
( 1 984) for no more than three weeks prior to tissue dissection. 



262 S. C. HAND 

Hyaluronidase (Type 1-S), collagenase (Type IV), DNAase I (Type IV), chymo- 
trypsin (Type II), soybean trypsin inhibitor, and Percoll were purchased from Sigma 
Chemical Co. NaH' 4 CO 3 was obtained from New England Nuclear. All other chemi- 
cals were reagent grade. Solutions of sodium sulfide were prepared fresh before each 
experiment (Powell and Somero, 1986a) and maintained under nitrogen until use 
(1-3 h). To reduce mechanical damage to isolated cells, siliconized glassware was 
used in all steps described below, and all pipets were firepolished. 

Tissue dissociation 

C. magnified gill tissue was placed on a chilled glass plate and minced into cubes 
varying in size from 0.5 mm to 2 mm. The tissue was rinsed briefly in Ca ++ -Mg ++ 
free salt solution (CMF solution) (508 mM NaCl, 10 mM KC1, 8.7 mM NaHCO 3 , 
28.6 mM Na 2 SO 4 , 0.1 mM EGTA, 4 mM glucose, pH 7.2) to remove mucus. The 
tissue was then transferred to 50 ml flasks and incubated for 1 5 min at 20C in 20 ml 
of CMF solution on a rotary shaker (1 10 cycles/min). This initial medium was re- 
placed with 10 ml of artificial seawater (4 1 1 mM NaCl, 9.6 mM KC1, 54 mM MgCl 2 , 
10.5 mMCaCl 2 , 8.8 mMNaHCO 3 , 23.6 mM Na 2 SO 4 , glucose 4 mM, pH 7.2) con- 
taining hyaluronidase (400 U/ml), collagenase (500 U/ml), and chymotrypsin (70 U/ 
ml), and the tissue was incubated for 1.5 h at 20C. At the end of this period, the 
tissue was rinsed with CMF solution and incubated 1 5 min in 10ml of CMF solution 
containing bovine serum albumin ( 1 mg/ml), trypsin inhibitor (0.4 mg/ml), and 
DNAase 1(15 U/ml). The tissue in this solution was flushed 20-30 times through a 
siliconized Pasteur pipet, a procedure that released large numbers of cells. The cellu- 
lar suspension was filtered sequentially through 250 micron and 100 micron nylon 
mesh (Tetko, Inc.; Elmsford, New York) to remove undissociated tissue. 

Isolated cells from Riftia trophosome were prepared similarly, except the tissue 
incubation with enzymes was shortened to one hour at 20C. The concentrations of 
enzymes were all reduced 50%, compared to the levels used for C. magnifica. 

The procedure for dissociation of gill tissue from L. floridana differed from the 
above protocol for C. magnifica in several respects. The concentration of the artificial 
seawater was 40 ppt (470 mM NaCl, 11 mM KC1, 62 mM MgCl 2 , 12mMCaCl 2 , 10 
mM NaHCO 3 , 27 mM Na 2 SO 4 , 0.1 mM EGTA, 5 mM glucose, pH 7.2), and the 
CMF solution consisted of 581 mM NaCl, 1 1.4 mM KC1, 10 mM NaHCO 3 , 32.7 
mM Na 2 SO 4 , 0.1 mM EGTA, 5 mM glucose, pH 7.2. The minced gill tissue was 
incubated for 15 min at 37C in CMF solution, and the concentrations of enzymes 
used in the subsequent incubation (1 h at 37C) were 50% of those used for C. mag- 
nifica tissue. 

Cell isolation 

The cellular suspension from C. magnifica gill was divided into two 5-ml portions, 
each of which was layered onto a Percoll gradient at 4C. This gradient separated 
bacteriocytes from other cell types and from acellular and subcellular debris. Cellular 
suspensions ofR. pachyptila trophosome were treated similarly. The 40-ml discontin- 
uous gradient consisted of four steps of 10% (density, 1.042), 30% (1.065), 50% 
(1.089), and 70% (1.111) Percoll. Each step was prepared by adding appropriate 
amounts of Percoll and deionized water to 2 ml of a concentrated CMF stock (5X). 
The cells settled without centrifugation for 3 h at 4C, and cells that had accumulated 
at each interface were collected and rinsed twice with artificial seawater to remove 
Percoll (which interferes with the assay for CO 2 incorporation). 



INVERTEBRATE-BACTERIA SYMBIOSES 263 

The Percoll gradient was changed to 30%, 50%, 70%, and 90% (density, 1 . 1 20) for 
separation of L. fioridana bacteriocytes. Each step was prepared in the 40 ppt CMF 
solution (final concentration). 

Cell concentrations were determined with a hemocytometer. The distinguishing 
features used to identify bacteriocytes under light microscopy were their granular 
appearance, lack of cilia, and relatively large diameter (20 microns, C. magnified; 40 
microns, L. fioridana). 

Transmission electron microscopy 

Isolated cells to be fixed for electron microscopy were transferred to Beem cap- 
sules and centrifuged at low speed (500 X g) to concentrate the cells. The supernatant 
was removed, and glutaraldehyde (4% in 0.3 M PIPES buffer, pH 7.2 at room temper- 
ature) was layered over the cells. Intact tissue for fixation was dissected into small 
blocks ( 1 mm diameter) and placed in plastic specimen trays containing glutaralde- 
hyde solution. After 30-60 min, the glutaraldehyde was removed, and the Beem cap- 
sule (or specimen tray) was filled with warmed agar (1.5% in 0.3 M PIPES, pH 7.2). 
After the agar solidified, the capsules were given three 15-min washes in buffer and 
then placed in 1% osmium tetroxide (prepared in 0.2 M potassium phosphate buffer, 
pH 7.4) for 2-3 h. The preparations were washed thoroughly with deionized water, 
dehydrated in a graded acetone series, and embedded in Spurr's low-viscosity media. 
All the steps above were completed on board ship. Sections were cut with a Sorvall 
MT 5000 Ultramicrotome and stained with 4% uranyl acetate followed by lead ci- 
trate. Cells were viewed with a Hitachi H-600 electron microscope. 

CO 2 fixation studies 

Isolated cells (100,000-500,000 for each assay) were incubated for up to 30 min 
in 0.5 ml of artificial seawater (pH 8.2) containing 1 microcurie of NaH 14 CO 3 , with 
and without various concentrations of sulfide and thiosulfate. All incubations were 
performed at 20C. Incorporation of CO 2 was stopped by vigorously mixing 0.1 ml 
of 1 2 N HC1 with each sample. Samples were transferred to plastic counting vials 
and heated for 2 h at 90C. Radioactivity remaining in the acid soluble fraction was 
quantified with liquid scintillation counting. Values for duplicate samples stopped at 
time were subtracted from all treatments. 

A TP measurements 

Experiments to determine the influence of sulfur compounds on cellular ATP 
levels were performed similarly to those above, but the radioactive bicarbonate was 
omitted from the incubation medium. At the end of the incubation, cells were sedi- 
mented with low speed centrifugation (500 X g, 5 min) at 4C, and the incubation 
medium was decanted. Cells were then resuspended in 0.5 ml of ice-cold 0.6 M per- 
chloric acid and homogenized. The homogenate was neutralized (and perchlorate 
salts precipitated) with 0. 1 5 ml of a solution containing 0.2 7VKOH, 0.4 Mimidazole, 
and 0.4 M KC1. The supernatant was initially stored in liquid nitrogen on board ship 
and later transferred to a -80C freezer until ATP analyses could be performed. 

ATP was measured with an enzyme-coupled fluorometric assay (modified from 
Lowry and Passonneau, 1972). All solutions were filtered through Gelman TCM-450 
(0.45 micron) filters before use. The 1 .2 ml assay mixture contained 50 mMTris-HCl 
buffer pH 8.1, 1.0 mM MgCl 2 , 0.2 mM dithiothreitol, 0.05 mM NADP, 0.1 mM 
glucose, and 50 microliters of sample. First, 0.07 units of glucose-6-phosphate dehy- 



264 S. C. HAND 

drogenase were added to eliminate endogenous G-6-P, and then 0.34 units of hexoki- 
nase were added for the quantification of ATP. The excitation wavelength was 365 
nm, and the emission monochromator was set at 460 nm. The increase in fluores- 
cence was measured with a Turner Model 430 spectrofluorometer, and the fluores- 
cence signal was adjusted so that 0. 1 nmole of ATP gave a 25% full scale deflection. 

Protein measurements 

Total protein was analyzed following the procedure of Peterson (1977). 

RESULTS 

Morphology oftrophosome tissue 

Fresh trophosome from R. pachyptila is a gelatinous, pulpy, dark iridescent-green 
tissue. If the tissue is suspended in saline, numerous finger-like lobules project into 
the medium producing a villous appearance. Each lobule is approximately 0. 1 5 mm 
in diameter and has a complex ultrastructure that is revealed by examining cross 
sections with electron microscopy (Fig. 1 A). 

The cells composing the outer pigmented layer of the lobule are tightly packed 
with membrane-bound inclusions of at least three morphological types. One type of 
granule (Fig. IB, 2A) is homogeneous in composition, weakly electron-dense, and 
similar in appearance to mucus droplets or mucigen granules found in goblet cells of 
intestinal epithelia (e.g.. Porter and Bonneville, 1973). In contrast, the darker osmio- 
philic granules (Fig. IB, 2B) contain highly organized, crystalline arrays of material 
(probably proteinaceous) that may be responsible for the intense green color of the 
trophosome. Indeed, if this outer cellular layer is osmotically lysed and removed from 
fresh trophosome tissue, the underlying tissue is white (R. Vetter, pers. comm.). The 
third type of inclusion (Fig. IB) has an electron density intermediate to that of the 
previous two granules, and its appearance indicates a heterogeneous composition. 
Thus, based on the internal morphology, these pigmented cells comprising the outer 
layer oftrophosome tissue are referred to hereafter as trophochrome cells. 

Nuclei are visible in the trophochrome cells, but other common organelles (e.g., 
mitochondria, endoplasmic reticulum) are infrequent. The granules described above 
occupy the vast majority of the intracellular space. Even though it is possible that 
osmotic swelling could have accentuated the size of these inclusions, the structural 
integrity of both the limiting and internal cell membranes does not suggest extensive 
swelling. 

Subtending the trophochrome cell layer are numerous bacterial endosymbionts 
(Fig. 3A, B). The bacteria housed in trophosome tissue are roughly spherical, often 
with irregular cell envelopes, and are approximately 3-5 microns in diameter. As one 
moves toward the center of the lobule, the morphology of the bacteria changes. Sulfur 
deposits within bacteria increase in both size and number, and ribosomes in the bac- 
terial cytoplasm are less distinct compared to bacteria located toward the periphery 
of the lobule (Fig. 3 A, B). (Note: sulfur deposits are identified as vacuoles where sulfur 
was extracted during tissue dehydration and embedding procedures.) Bacteria located 
toward the center of the lobule also appear smaller. Concentric membrane whorls 
within the bacteria are occasionally visible (Fig. 4 A, upper left). 

Although it is difficult to fully trace eucaryotic cell membranes, the following evi- 
dence indicates that the vacuole-enclosed bacteria are located within animal cells 
(bacteriocytes). There are nuclei, mitochondria, and other organelles interspersed 
among the bacteria (Fig. 4A, B). In Figure 4A, it is possible to delineate (moving 



INVERTEBRATE-BACTERIA SYMBIOSES 



265 





FIGURE 1 . A. Cross section of a trophosome lobule from Riftia pachyptila viewed at low magnifica- 
tion with transmission EM. The outer trophochrome cell layer contacts the coelomic fluid space (cf). 
Subtending this pigmented cell layer are numerous bacteria (b). At the center of the lobule is a hemolymph 
space (hs). Scale bar = 10 microns. B. Higher magnification of the trophochrome cells showing a nucleus 
(n) and the tight intracellular packaging of diverse types of granules. Scale bar = 3 microns. 



266 



S. C. HAND 



\ 









FIGURE 2. Two types of vacuole-enclosed granules present in trophochrome cells from Riftia pa- 
c/iyptila. A. Nondescript granules of uniform density, similar in appearance to mucigen granules. Scale bar 
= 1 micron. B. Electron-dense granule containing crystalline arrays (ca) of material that is proteinaceous in 
appearance. Scale bar = 2 microns. 



outward from the center of the bacterium labeled "b") the bacterial cell envelope, the 
peribacterial membrane, and immediately adjacent, a nuclear envelope. 

At the very center of the trophosome lobule, there is a hemolymph space or sinus 
extending longitudinally (Fig. 4B). We were unable to discern a basal lamina separat- 
ing the bacteriocytes from the hemolymph space of the lobule. 

Since the morphology and fine structure of the gill tissue from C. magnified (Fiala- 
Medioni and Metivier, 1986) and L. floridana (Fisher and Hand, 1984) have already 
been described, we will not redescribe them here. 



Isolated cell preparations 

The cell purity for bacteriocytes isolated from bivalve gill tissue was approxi- 
mately 70-80 percent (Table I). The amount of cellular debris present was not quanti- 
fied, but was generally low. The yield of bacteriocytes was higher from L. floridana 
gill tissue than from C. magnified. The reason may be that, while dissociation of L. 
floridana tissue was performed at 37C, the incubation temperature for C. magnifica 
tissue had to be reduced to 20C because of its temperature sensitivity. Although 
using 37C incubations with the vent clam tissue improved the yield, the resulting 
bacteriocytes were not viable, as judged by the lack of ability to fix CO 2 . Prior to the 
Percoll gradient step, the bacteriocytes represented 1 1% of the total cells in suspension 
from L. floridana gill. Thus, the density gradient fractionation achieved a 7-fold en- 
richment of bacteriocytes. 

Attempts to isolate intact bacteriocytes from trophosome tissue were unsuccess- 
ful, suggesting that these cells are very fragile and are unable to withstand the isolation 



INVERTEBRATE-BACTERIA SYMBIOSES 



267 






f.Vf. J * 

: *V 

*- 






' . 

1v-.';-;.- 





I " - 





FIGURE 3. Bacteria (b) of Riftia pachyptila trophosome tissue. A. Located toward the periphery of 
the lobule, these bacteria are granular and contain very few sulfur deposits (observed as holes in the section 
where leaching has occurred). Scale bar = 1 micron. B. Bacteria located toward the center of the lobule 
contain more sulfur vacuoles (v), and their cytoplasm is less granular. Scale bar = 1 micron. 



268 



S. C. HAND 





FIGURE 4. A. Bacterium (b) immediately adjacent to an animal cell nucleus (n). Scale bar = 1 mi- 
cron. B. Micrograph showing hemolymph spaces (hs) located at the center of the lobule. Scale bar = 5 
microns. 

procedures. In contrast, trophochrome cells were enriched to a purity of 80%, and 
free bacteria were isolated in high yield and at a comparable purity to the pigmented 
trophosome cells. The primary contaminants of the trophochrome cell preparation 
were free bacteria. 



INVERTEBRATE-BACTERIA SYMBIOSES 269 

TABLE I 



Yield and purity of isolated cell preparations 



Gradient interface Yield Purity Protein 
Cell source and type (% Percoll) (million cells/g tissue) (% of total) (mg/million cells) 

C. magnified 

Bacteriocytes 1.160.08 a 72 5.6 0.90 0.1 4 

30-50%, 50-70% (n = 3) (n = 5) (n = 3) 
Non-bacteriocyte 

epithelial cells 10-30% 10.3 97 0.16 

L. floridana 



Bacteriocytes 


50-70% 


3.0 81 0.8 
(n = 3) 





R. paehyptila 
Bacteria 


10-30% 


79, 70 


0.014 


Trophochrome cells 


30-50%, 50-70% 


78,82 


1.09 



a Mean standard error. 



One prominent ultrastructural feature of isolated bacteriocytes from C. magnified 
is wide-spread fields of bacteria easily the most numerous subcellular structures in 
the cytoplasm (Figs. 5A, C; 6A). Compared to the bacteria present in trophosome 
tissue, these bacteria are much smaller (0.5-0.7 micron diameter). They are clearly 
contained in vacuoles within the bacteriocyte, and nuclear regions are evident 
(Fig. 6A). 

Large granules (approximately 5- 1 micron diameter) are a second salient feature 
of bacteriocytes from both C. magnified (Fig. 5 A, C) and L. floridana (Fig. 5B). Some 
of the C magnified granules are irregularly shaped and quite electron-dense, and 
others are more circular and have a stippled appearance. The electron-dense granules 
are morphologically similar to those of bacteriocytes from L. floridana (Fig. 5B, and 
Fisher and Hand, 1984). When viewed with Nomarski differential-interference-con- 
trast microscopy, the granules of L. Floridana are striking and are certainly the domi- 
nant inclusion of the bacteriocyte (Fig. 5B). Previous work indicated the presence of 
iron in these granules (Fisher and Hand, 1984; cf., Wittenberg, 1985), but whether 
the bacteriocytes' granules in C. magnified are chemically similar is unknown. As 
judged by the sedimentation behavior of bacteriocyte populations in Percoll gradi- 
ents, L. floridana bacteriocytes have a greater density, which may be a consequence 
of differences between these pigment granules. Bacteriocytes from L. floridana also 
are over two-fold larger in diameter than those from C. magnified (Fig. 5A, B). 

The morphology of isolated trophochrome cells (Fig. 6B) is essentially unchanged 
from that seen for intact tissue (Fig. IB). On the other hand, isolated bacteria look 
more irregular in overall shape compared to those viewed in situ and occasionally 
were more vacuolated, suggesting that the isolation procedure for this cell type needs 
improvement. 

Metabolic properties of isolated cells 

The capacity for carbon dioxide fixation of C. magnified bacteriocytes and iso- 
lated bacteria from Rift id trophosome is presented in Table II. In the absence of sulfur 
compounds, incorporation of CO 2 by bacteriocytes proceeds at a rate of 13 nmoles 
CO 2 /mg protein/h. Fixation is inhibited by sulfide and, to a lesser degree, thiosulfate 



270 



S. C. HAND 





FIGURE 5. A. Isolated bacteriocytes from the gill tissue of Calyptogena magnified viewed with trans- 
mission EM. Two distinctly different granules (g) are observable in the cytoplasm. Scale bar = 1 5 microns. 
B. Nomarski light micrograph of bacteriocytes from Lucinafloridana gill tissue, emphasizing the promi- 
nent granules in these very large cells. Scale bar = 60 microns. C. Higher magnification of C. magnified 
bacteriocytes illustrating the expansive fields of symbiotic bacteria (b). Nucleus (n), granules (g). Scale bar 
= 5 microns. 



INVERTEBRATE-BACTERIA SYMBIOSES 



271 







FIGURE 6. A. High magnification (78,000 x) of the symbiotic bacteria present in the cytoplasm of 
isolated bacteriocytes from Calyptogena magnified. Note the peribacterial membrane encompassing each 
bacterium, and the distinct nuclear regions. The bacteria are approximately an order of magnitude smaller 
than those in Riftia trophosome. Scale bar = 0.5 micron. B. Isolated trophochrome cell from Riftia pachyp- 
tila. Scale bar = 5 microns. 



across the range of concentrations used here. The rate of fixation by L. floridana 
bacteriocytes is approximately doubled by the addition of 0. 1 mM thiosulfate; higher 
thiosulfate concentrations do not appreciably alter the fixation rate. Incorporation 
by non-bacteriocyte epithelial cells of C. magnified is approximately one third the 
rate of C. magnified bacteriocytes. The incorporation by the non-bacteriocyte prepa- 
ration is presumably due to eucaryotic enzymes like pyruvate carboxylase and phos- 
phoenolpyruvate carboxykinase, both known to occur in marine bivalves (e.g., Fel- 
beck, 1983; Meinardus-Hagar and Gade, 1986). Contamination by bacteriocytes in 
this preparation is low (Table I). 

The positive influence of sulfide on the intracellular ATP levels of C. magnified 
bacteriocytes contrasts with the inhibitory effect seen on CO 2 fixation (Table III). 
After 60 min, bacteriocytes incubated in 0.5 mM sulfide contain 3.30 nmoles ATP/ 
million cells (approximately 1 .0 1 mM) , compared to 1 .02 nmoles/million cells (0.3 1 
mM) with no added sulfur, and 1.74 nmoles/million cells (0.54 mM) with 0.5 mM 
thiosulfate. 



DISCUSSION 

The primary objectives of this study were ( 1 ) to describe ultrastructural features of 
Riftia trophosome tissue, particularly those related to the distribution and subcellular 
location of bacteria and the nature of the pigmented layer of trophochrome cells, (2) 
to prepare suspensions of isolated cells from invertebrate tissues that contain bacterial 
endosymbionts, and (3) to use these cellular preparations for characterizing meta- 
bolic aspects of the symbioses. 

Morphological evidence supports the conclusion that symbiotic bacteria present 
in the trophosome of R. pachyptila are contained within animal cells (i.e., bacterio- 



272 S. C. HAND 

TABLE II 

Incorporation of carbon dioxide in isolated celt suspensions at 20C 



Preparation 


Sulfur compound 
present 


CPM/million 
cells/h 


n moles CO 2 / 
million cells/h 


nmoles CO 2 / 
mg protein/h 


C. magnified 
Bacteriocytes 


none 


5632 
5976 


13.9 

14.8 


13.0 
13.3 




Na 2 S: 0. 1 mM 


4590 


11.4 









4272 


10.6 


9.5 




0.5 mM 


3696 


9.1 









4224 


10.5 


9.5 




1 .0 mM 


2676 


6.6 







Na 2 S 2 O 3 :0.1 mM 


4488 
5904 


11.1 
14.6 


13.2 




0.5 mM 


3084 


7.6 








3684 


9.1 


8.1 




1.0 mM 


3792 


9.4 


8.4 


Non-bacteriocyte 
epithelial cells 


none 
Na 2 S: 0. 1 mM 
0.5 mM 


292 
404 
302 


0.72 
0.10 
0.75 


4.5 
6.2 
4.6 




1.0 mM 


318 


0.79 


4.8 




1.5 mM 


348 


0.86 


5.3 




2.0 mM 


136 


0.34 


2.1 




Na.S.OvO.l mM 
0.5 mM 


292 
266 


0.72 
0.66 


4.5 
4.0 




1.0 mM 


246 


0.61 


3.8 




1.5mA/ 


274 


0.68 


4.2 




2.0mM 


292 


0.72 


2.2 


L. floridana 
Bacteriocytes 


none 
Na 2 S 2 O,:0.1 mM 
0.2 mM 


3440 

7776 
7176 


8.4 
19.0 
17.6 







0.5 mM 


8728 


21.4 







0.75 mM 


7912 


19.4 







1.0 mM 


8824 


21.6 





R. pachyptila 
free bacteria 


none 








13.5 



cytes). Electron micrographs show that nuclei and other eucaryotic organelles are 
frequently interspersed among the vacuole-enclosed bacteria, and eucaryotic cell 
membranes appear to enclose bacteria and such organelles within the same cell. Cava- 
naugh (1985) suggested that an intracellular location for the bacteria was probable, 
but previous evidence has been inconclusive (Cavanaugh et al, 1981; Cavanaugh 
1983, 1985). 

A second notable point regarding trophosome fine structure is the distribution of 
sulfur deposits among the bacteria within a trophosome lobule. The bacteria located 
closer to the outer trophochrome layer clearly have fewer deposits than do bacteria 
located nearer the hemolymph space at the center of the lobule. This spatial distribu- 
tion could reflect greater access of the latter bacteria to high sulfide concentrations in 



INVERTEBRATE-BACTERIA SYMBIOSES 

TABLE III 

Intracellular A TP levels of Calyptogena magninca hacteriocvtes in the presence 
of sulfur compounds at 20C 



273 









Calculated intracellular 


Incubation time 1 


Sulfur 


nmoles ATP/ 


ATP concentration 


(min) 


compound 


million cells 


(mM) 2 





none 


1.34 


0.41 


30 


none 


0.83 


0.25 




0.5mAfNa 2 S 


1.34 


0.41 




0.5mA/Na 2 S 2 O;, 


0.88 


0.28 


60 


none 


1.02 


0.31 




0.5mA/Na 2 S 


3.30 


1.01 




0.5mA/Na 2 S 2 O 3 


1.74 


0.54 


90 


none 


2.03 


0.62 




0.5mMNa 2 S 


3.75 


1.15 




0.5 mAl Na 2 S 2 O 3 


1.29 


0.40 



'Cells were incubated at 20C in artificial seawater (pH 8.2) containing the indicated concentrations 
of sulfide and thiosulfate. 

2 The intracellular concentrations of ATP were calculated using an average bacteriocyte diameter of 
19.8 0.8 microns (SE, n = 25), as determined from transmission electron micrographs. Cellular water 
content was assumed to be 80%. 



Rift ia vascular blood (Childress et ai, 1984). Indeed, Vetter (1985) showed that in 
the absence of sulfide, sulfur globules are lost from bacterial symbionts in the gill 
tissue of Lucinoma anmdata; presumably the presence of sulfide would stimulate 
deposition of these elemental sulfur stores. The occurrence and possible roles of sulfur 
deposits in free-living sulfur bacteria recently have been reviewed (Vetter, 1985). It 
should be noted that the coelomic fluid ofRiftia also contains sulfide, but it is unclear 
whether this sulfide pool is available directly to the symbionts, or must first be trans- 
ferred to the vascular circulation (Childress et ai, 1 984). If the density of sulfur depos- 
its is related to the level of sulfur-based metabolism occurring in a given bacterium, 
then a gradient of metabolic activity could exist, with higher metabolic potential be- 
ing possessed by bacteria closer to the central hemolymph space of the lobule. Bacte- 
ria near the hemolymph space also appear smaller, which is consistent with them 
being younger than the larger bacteria toward the outer trophochrome layer. 

The morphology of the pigmented layer of trophosome tissue was far more com- 
plex than originally anticipated. First, the trophochrome layer does not contain bacte- 
rial endosymbionts, although the cells do contain extensive numbers of cytoplasmic 
inclusions (Figs. 1 B, 2A, B). The function(s) of these cells is not understood presently, 
but their internal structure suggests several possibilities. Some of the intracellular 
droplets or granules are structurally similar to those found in mucus-secreting cells 
(e.g., intestinal goblet cells). What benefit the secretion of mucus-like material (if it 
were to occur) by trophochrome cells would confer to the tubeworm is unclear, other 
than possibly lubricating the external surface of the trophosome that is in direct appo- 
sition to the internal surface of the body wall of the worm. Another possible function 
of trophochrome cells, that of phagocytosis and degradation of aging bacteria, is sug- 
gested by the heterogeneous contents of other intracellular granules (Fig. 1 B). Future 
experiments measuring levels of protease activity in isolated trophochrome cells 
could be enlightening in this context. Finally, trophochrome cells also contain elec- 
tron-dense material that is deposited as crystalline arrays in some granules (Fig. 4B). 



274 S. C. HAND 

This material is likely proteinaceous and may be the pigment responsible for the deep 
green color of the trophosome. 

The region of the trophosome containing the bacteriocytes is a diffuse, loosely 
associated tissue, which suggests that its dissociation into suspensions of single bacte- 
riocytes should have been relatively straight forward. Apparently however, the cells 
are very fragile, and we were unable to obtain intact bacteriocytes from trophosome 
even when mechanical agitation was kept to a minimum. 

Thus, the isolated cellular preparation that may have the most potential for im- 
proving our understanding of the mechanisms involved in metabolic utilization of 
sulfur by the symbiosis is the bacteriocytes from C. magnified and L. floridana. The 
ultrastructure of these isolated cells is comparable to the structure as viewed in situ 
(Fisher and Hand, 1984; Fiala-Medioni and Metivier, 1986), with the exception that 
bacteria in isolated bacteriocytes from C magnified appear a little more spherical 
than those in intact tissue (pers. obs., and Fiala-Medioni and Metivier, 1986). The 
size of C magnifica bacteria differs markedly from those of Rifiia trophosome, and 
data from 5S rRNA sequencing suggest the bacteria from the two sources have mini- 
mal affiliation (Stahl et ai, 1984; Lane et ai, 1985). 

The temperature sensitivity of C. magnifica bacteriocytes closely parallels the ob- 
servations of Belkin et ai (1986) in their studies with gill homogenates of the vent 
mussel Bathymodiolus thermophilus. In both cases, the capacity for CO 2 fixation de- 
clines dramatically at temperatures above 20C. Thus, the bacterial endosymbionts 
from C. magnifica are comparable to those of B. thermophilus in that both appear to 
be sulfur-oxidizing bacteria with psychrophilic characteristics (cf., Belkin et al, 1 986). 
The detrimental effect of warm temperature was not observed for bacteriocytes from 
the shallow-water bivalve L. floridana, and in general this cellular preparation seemed 
hardier. 

In the absence of an exogenously added sulfur source, the rate of CO 2 fixation by 
C. magnifica bacteriocytes was 1 3 nmoles/mg protein/h. Rates obtained with homog- 
enates of B. thermophilus gill ranged between 1.7 and 9.4 nmoles/mg protein/h, de- 
pending on temperature and the thiosulfate concentrations present (Belkin et al., 
1986). Using "purified" bacteria from B. thermophilus, Belkin et al, (1986) reported 
maximum CO 2 fixation rates of approximately 40 nmoles/mg protein/h in the pres- 
ence of thiosulfate. In the present study, CO 2 fixation by C. magnifica bacteriocytes 
was inhibited by sulfide across the range of 0.1 to 1.0 mM. Less inhibition was seen 
with thiosulfate. Similarly, Anderson (1986) showed that CO 2 fixation by whole speci- 
mens ofSolemya reidi was inhibited at sulfide concentrations above 0. 1 mM in sur- 
rounding seawater. This protobranch bivalve has a chemoautotrophic metabolism 
based on the presence of sulfur bacteria localized in its gill tissue (Felbeck, 1983). 
Unfortunately, we do not have measurements of CO 2 fixation by bacteriocytes at 
sulfide concentrations below 0.1 mM. Considering Anderson's results (1986), such 
sulfide levels could well stimulate the process. 

CO 2 fixation by L. floridana bacteriocytes was increased approximately two-fold 
by thiosulfate concentrations between 0. 1 and 1 .0 mM, a pattern virtually identical 
to that seen for gill homogenates of B. thermophilus (Belkin et al., 1 986). Exogenously 
added sulfide was minimally effective in stimulating the process in L. floridana bac- 
teriocytes (preliminary observations) and in B. thermophilus homogenates (Belkin et 
ai, 1986). The rate of CO 2 fixation by isolated bacteria from R. pachyptila tropho- 
some was 13.5 nmoles/mg protein/h in the absence of added sulfur. The response to 
sulfide and thiosulfate was highly variable, and thus the results are not reported here. 
This variability could be related to the general instability of the CO 2 fixation capacity 
exhibited by trophosome preparations (Belkin et ai, 1986). 



INVERTEBRATE-BACTERIA SYMBIOSES 275 

Although sulfide (0. 1 mA/and above) inhibited CO 2 fixation by isolated bacterio- 
cytes from C. magnified, the presence of 0.5 mM sulfide promoted higher intracellu- 
lar ATP concentrations than control incubations without any added sulfur. The op- 
posing effects of sulfide on the two processes may be related to the sulfide concentra- 
tions used; as discussed earlier, if lower sulfide levels had been tried, CO 2 fixation 
might also have been stimulated. Thiosulfate was not as effective in enhancing intra- 
cellular ATP levels. Powell and Somero (1986a), using lysed bacterial preparations 
from C. magnified gill, showed that sulfite stimulated ATP synthesis while thiosulfate 
and sulfide did not. In their study, ATP was measured using the firefly luciferase 
technique, and incubations were of shorter duration. Most likely, components of the 
animal cell contributed to sulfide utilization in our study with isolated bacteriocytes. 
Although some degree of spontaneous oxidation is possible, Powell and Somero 
(1985) localized sulfide oxidizing bodies in the animal cell cytoplasm of S. reidi gill 
tissue. These workers also have conclusively demonstrated that mitochondria iso- 
lated from Solemya reidi gill tissue can couple sulfide oxidation to the formation of 
ATP via the electron transport system and oxidative phosphorylation (Powell and 
Somero, 1986b). 

Thus, the initial step(s) of sulfide oxidation could occur in bacteriocytes prior to 
the sulfur compound reaching the endosymbiont, and some of the ATP synthesis in 
bacteriocytes could reflect mitochondrial processing. Consequently, the identities of 
sulfur compounds exploited for chemical energy by the symbiosis may depend on 
the structural organization and integrity of the biological preparation under study. 
Additional metabolic characteristics of invertebrate-sulfur bacteria symbioses may 
display similar dependencies in ways yet to be identified. 

ACKNOWLEDGMENTS 

Appreciation is extended to the captains and crew members of the R/V Melville, 
R/V Atlantis II, and DSRV Alvin. The expedition was supported by NSF grant 
OCE83- 1 1 256 (facilities support grant for the Galapagos '85 program; Drs. J. J. Chil- 
dress and K. L. Johnson, University of California, Santa Barbara, co-principal inves- 
tigators). Travel funds were supplied to SCH by the University of Southwestern Loui- 
siana. Helpful discussions of these findings with Drs. J. Pickett-Heaps and L. A. 
Staehelin (MCD Biology, University of Colorado) and Dr. S. Schmidt (EPO Biology) 
are gratefully acknowledged. Technical assistance was provided by Mr. V. Bullman, 
Mr. M. Fisher and Mr. L. Harwood, and advice concerning the fixation and embed- 
ding procedures for isolated cells was provided by Drs. R. C. Brown and B. E. Lem- 
mon (University of Southwestern Louisiana). 

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Galapagos rift hydrothermal vents. NATO Conf Ser. (Afar. Sci.) 12: 735-770. 
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symbionts by ribosomal RNA sequences. Bull. Biol. Sot: Wash. 1985(6): 389-400. 
LOWRY, O. H., AND J. V. PASSONNEAU. 1972. A Flexible System of Enzymatic Analysis. Academic Press, 

New York. 291 pp. 

MEINARDUS-HAGER, G., AND G. GADE. 1 986. The pyruvate branchpoint in the anaerobic energy metabo- 
lism of the jumping cockle Cardium tuherculatum L.: D-lactate formation during environmental 

anaerobiosis versus octopine formation during exercise. Exp. Biol. 45: 91-110. 
PETERSON, G. L. 1977. A simplification of the protein assay method of Lowry et al. which is generally 

more applicable. Anal. Biochem. 83: 346-356. 
PORTER, K. R., AND M. A. BONNEVILLE. 1973. Fine Structure of Cells and Tissues. Lea and Febiger, 

Philadelphia. 204 pp. 
POWELL, M. A., AND G. N. SOMERO. 1983. Blood components prevent sulfide poisoning of respiration of 

the hydrothermal vent tube worm Riftia pachyptila. Science 219: 297-299. 
POWELL, M. A., AND G. N. SOMERO. 1985. Sulfide oxidation occurs in the animal tissue of the gutless 

clam, Solemya reidi. Biol. Bull. 169: 164-181. 
POWELL, M. A., AND G. N. SOMERO. 1986a. Adaptations to sulfide by hydrothermal vent animals: sites 

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POWELL, M. A., AND G. N. SOMERO. 1 986b. Hydrogen sulfide oxidation is coupled to oxidative phosphor- 

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Reference: Biol. Bull. 173: 277-288. (August, 1987) 



ENERGETICS OF CONTRACTILE ACTIVITY IN ISOLATED RADULA 

PROTRACTOR MUSCLES OF THE WHELK BUSYCON CONTRARIUM: 

ANAEROBIC END PRODUCT ACCUMULATION AND RELEASE 

ROBERT W. WISEMAN AND W. ROSS ELLINGTON 1 

Department of Biological Science, The Florida State University. Tallahassee, Florida 32306-3050 

ABSTRACT 

Anaerobic energy metabolism during contractile activity was investigated in the 
isolated radula protractor muscle of the whelk Busycon contrarium. Spectrophoto- 
metric assay of enzyme activities in crude tissue extracts revealed particularly high 
pyruvate reductase activities with octopine dehydrogenase displaying the highest ac- 
tivity. During electrically induced isotonic contractions of the radula protractor mus- 
cles, the following end products, listed in order of increasing level, accumulated in 
the tissue: strombine, octopine and alanopine (the "opines"), and D-lactate. Pyruvate 
levels increased three-fold during muscle contraction, suggesting that pyruvate plays 
a key role in the regulation of the pyruvate reductases. The muscle released lactate, 
but none of the opines, into the incubation medium, with rates exceeding 3 ^moles 
min ' -g wet wt '. During the later phases of contraction, more D-lactate was re- 
leased into the medium than accumulated in the muscle. We conclude that transport 
of D-lactate permits sustained flux through lactate dehydrogenase because of a reduc- 
tion in product inhibition. Furthermore, we hypothesize that D-lactate transport may 
be coupled to H + export or OH import, which would then serve to regulate the 
extent of accumulation of glycolytically produced protons. 

INTRODUCTION 

The muscles of marine molluscs possess two distinctly different mechanisms of 
energy production during periods of reduced oxygen availabiliy. During environmen- 
tal anaerobiosis i.e., whole-organism exposure to anoxia aspartate and glycogen 
are cofermented, yielding succinate and alanine as end products (Gade, 1983). The 
succinate pathway typically occurs at relatively low rates and is associated with a 
reverse Pasteur effect in these muscles (Storey, 1985). A number of molluscs may 
undergo functional anaerobiosis, where oxygen demand exceeds delivery. In this in- 
stance, only certain tissues are rendered anoxic (Gade, 1 983). Under these conditions, 
the glycolytic flux is several orders of magnitude higher than under conditions of 
environmental anaerobiosis (Livingstone, 1982; Gade, 1983). The higher energy out- 
puts necessary for burst activity are provided by glycogen fermentation and the shunt- 
ing of pyruvate through pyruvate reductases such as lactate and opine dehydroge- 
nases (Gade and Grieshaber, 1 986), resulting in the accumulation of D-lactate, octo- 
pine, alanopine, or strombine. 

Opine dehydrogenases catalyze the reductive condensation of pyruvate and an 
amino acid according to the following general reactions: 



Received 22 September 1986; accepted 18 May 1987. 
' To whom reprint requests should be sent. 



277 



278 R. W. WISEMAN AND W. R. ELLINGTON 

arginine + pyruvate + NADH ^ octopine + NAD 

(ODH, octopine dehydrogenase) 

alanine + pyruvate + NADH ^ alanopine + NAD 

(ADH, alanopine dehydrogenase) 

glycine + pyruvate + NADH ^ strombine + NAD 

(SDH, strombine dehydrogenase) 

Many molluscan muscles have the enzymatic potential for producing several opine 
end products as well as D-lactate since high activities of opine and D-lactate dehy- 
drogenases may occur in the same tissue (Zammit and Newsholme, 1976; Living- 
stone el al., 1983). For example, the pedal retractor (Baldwin et al, 1981; Baldwin 
and England, 1982) and radula (Ellington, 1982) muscles of gastropods contain sig- 
nificant activities of all opine dehydrogenases as well as D-lactate dehydrogenase. The 
relative contribution of these enzymes to the maintenance of glycolytic flux during 
contractile activity, not yet fully explored, is considered in this paper. 

The metabolic disposition of end products of anaerobic metabolism in molluscs 
(Ellington, 1983b) is poorly known. Propionate is released into the hemolymph of 
Mytilus edulis (Zurburg et al., 1982), whereas succinate appears to accumulate in 
the hemolymph of the clam Mercenaria mercenaria (Korycan and Storey, 1983). 
Octopine is not released into the hemolymph during contractile activity in the giant 
scallop, Placopecten magellicanus (de Zwaan et al., 1980). However, hemolymph 
octopine levels are slightly elevated after contractile activity or hypoxia in the cepha- 
lopods Sepia officinalis and Loligo vulgaris (Storey and Storey, 1979; Gade, 1980). 
Alanopine and strombine appear not to be released from molluscan muscles. Octo- 
pine, alanopine, and strombine levels fall during recovery, indicating oxidation in 
situ (Ellington, 1983b). 

The present study focuses on the metabolism of the radula protractor muscle of 
the large marine gastropod Busycon contrarium. This muscle possesses relatively high 
activities of lactate and opine dehydrogenases. The presence of several pyruvate re- 
ductases in the radula protractor muscle poses questions about the control of these 
enzymes and the disposition of their products. We show that electrical stimulation of 
this muscle while it is subjected to anoxia induces the formation of all opines as well 
as D-lactate. Interestingly, formation of D-lactate is much greater than that of the 
other end products, even though the activities of the opine dehydrogenases are much 
higher. Further, D-lactate is released from the exercising muscle into the medium 
while the opines are retained. End-product removal may enhance the formation of 
additional D-lactate and, as a result, large amounts of carbon can be shunted through 
lactate dehydrogenase allowing for higher, sustained glycolytic fluxes during anoxia. 

MATERIALS AND METHODS 
Animals 

Specimens of the whelk Busycon contrarium were collected off Alligator Point in 
Franklin County, Florida, and were maintained in the flowing-seawater system at the 
Florida State University Marine Laboratory near St. Theresa. Individuals used in 
experiments were transferred to the Florida State University campus, where they were 
maintained for brief periods in a recirculating seawater system. 



ANAEROBIC METABOLISM OF RADULA MUSCLES 279 



Biochemicals 



Biochemicals were purchased from Boehringer-Mannheim (Indianapolis) and 
Sigma Chemical Company (St. Louis). D-Lactate dehydrogenase, used to determine 
D-lactate, was purified from the muscle of the horseshoe crab Limulus polyphemus. 
Octopine dehydrogenase, used to assay for octopine, arginine, and arginine phos- 
phate, was purified from the adductor muscles of the scallop Argopecten irradians 
concentricus. Succinyl Co A synthase, used in succinate assays, was a gift from Dr. 
William Bridger, Department of Biochemistry, University of Alberta, Edmonton. 

Experimental procedure 

Intact radula protractor muscles, dissected from the proboscis apparatus, were 
ligated at both ends with surgical silk and placed in a 5 X 75-mm muscle bath filled 
with 1.5 ml of MBL (Marine Biological Laboratory) formula artificial seawater 
buffered with 5 mA/hydroxyethylpiperazine ethanesulfonic acid (pH = 7.8). One end 
of the muscle was fastened to a hook electrode and pulled into a rubber sleeve at the 
bottom of the muscle bath. The other end was attached to a Narco Biosystems iso- 
tonic myograph transducer with the silk suture. Muscles were suspended in the bath 
at 1.5 times their resting length (measured upon excision). Temperature was main- 
tained by immersion of the bath in a larger water-filled vessel which was jacketed and 
controlled by a Brinkmann model RM 6 recirculating water bath (20C). Contrac- 
tions were recorded with the isotonic transducer connected to a Narco Biosystems 
model MK IV physiograph. A second electrode, inserted in the bath, delivered 
square-wave pulses (60 volts, 40 ms) at 2.5-s intervals from a Grass model SD9 stimu- 
lator. The bath was gassed with normocapnic nitrogen (0.05% CO 2 ) through a 75- 
mm, 22-gauge Luer lock syringe needle. 

Each muscle preparation was fastened in the bath and bubbled with nitrogen for 
1 5 min. Control experiments were terminated at this point, and the tissues were re- 
moved, blotted, and frozen in liquid nitrogen. Experimental groups were gassed as 
the controls were, and then subjected to various periods of electrical stimulation (2.5, 
5, 10, and 15 min) in the presence of normocapnic nitrogen before being frozen. 
In all cases, the medium was decanted from the bath and stored at -70C for later 
analysis. 

Enzyme assays 

Freshly dissected muscles were homogenized in 24 volumes of extraction buffer 
(50 mA/triethanolamine, pH 7.4, 1 mA/EDTA, 20 mM mercaptoethanol, 20% glyc- 
erol) with a Tekmar UltraTurrax tissue homogenizer. The homogenate was centri- 
fuged at 1 2,000 X g for 20 min at 4C. The supernatant was passed through a Sepha- 
dex G-25 column (1.5 X 14 cm) equilibrated with extraction buffer less glycerol, 
which removed low-molecular-weight compounds. The proteins in the void volume 
were used as the source of enzyme activities. The activities were determined spectro- 
photometrically with a Gilford model 252-1 spectrophotometer according to the 
methods outlined by Ellington (1982). 

Metabolite assays 

Nuetralized perchloric acid extracts were prepared from the radula protractor 
muscles frozen at -70C according to the methods of Graham and Ellington (1985). 
Arginine phosphate was assayed spectrophotometrically by the method of Grieshaber 



280 R. W. WISEMAN AND W. R. ELLINGTON 

TABLE I 

Profile ofpvruvate reductase activities in desalted tissue extracts ofradula protractor muscles from 
Busycon contrarium 

Enzyme Activity 



D-Lactate dehydrogenase 36.46 9.7 1 

Strombine dehydrogenase 6 1 .98 1 4.88 

Alanopine dehydrogenase 96.87 19.66 

Octopine dehydrogenase 509.83 40.73 

Activities are expressed in //moles min" ' g wet wt~ ' and were measured at 25C. Data represents mean 
1 SD, n = 4. 



and Gade (1976). Pyruvate was assayed fluorometrically in a Farrand Optical model 
A-4 fluorometer by the method of Lowry and Passonneau (1972). Both pyruvate 
and arginine phosphate levels were determined immediately after neutralization to 
eliminate sample loss. 

Aspartate, succinate, arginine, and malate were determined spectrophotometri- 
cally according to the method of Williamson and Corkey ( 1 969), Williamson ( 1 974), 
Grieshaber and Gade ( 1 976), and Williamson and Corkey ( 1 969), respectively. Octo- 
pine and lactate were assayed fluorometrically essentially as outlined by Graham and 
Ellington (1985). 

Concentrations of the free amino acids alanine and glycine were determined by 
HPLC on a Dionex amino acid analyzer with a Pierce amino acid column and buffers 
(Pierce Chemical Company). Alanopine and strombine concentrations were also de- 
termined by HPLC methods (Fiore el ai, 1984). 

RESULTS 
Enzyme activities 

Freshly prepared extracts of the radula protractor muscles of Busycon contrarium 
displayed high activities of all four pyruvate reductases (Table I). Octopine dehy- 
drogenase (ODH) had the highest activity of the enzymes assayed. Alanopine dehy- 
drogenase (ADH) displayed somewhat lower activity followed by strombine (SDH) 
and D-lactate (LDH) dehydrogenases (Table I). 

Contractile activity 

Contractile activity under nearly anoxic conditions was maintained within 98% 
of initial values for the first 5 min of electrical stimulation (Fig. 1 ). There was a general 
trend towards a decline in force thereafter. This pattern was evident in all muscle 
preparations tested. 

Metabolite levels in the tissue and the medium 

Arginine phosphate levels declined at the onset of muscular activity, reached a 
minimum after 10 min, and remained relatively constant thereafter (Table II). Free 
arginine levels increased in the first 2.5 min of contractile activity, then fell to near 
control levels at the end of the experiment (Table II). Within 10 min, alanine levels 
were three times greater than levels measured initially, but returned to near control 
levels after 15 min (Fig. 2). Glycine levels did not change significantly during the 15 



ANAEROBIC METABOLISM OF RADULA MUSCLES 



281 



100. 
80. 

0) 

60 J 
g 

I 40 - 

2? 

20 . 




10 



15 



mm 



FIGURE 1. Changes in contractile force for electrically stimulated isotonic contractions of isolated 
radula protractor muscles from Busycon contrarium versus time. Units are percent change of initial force. 
Points represent means 1 SD, n = 5. 



min of contractile activity (Fig. 2). Aspartate levels in the muscle declined during 
contractile activity, with the bulk of the change taking place in the first 5 min (Fig. 
2). Succinate and malate levels in muscles stimulated for 15 min were less than 0.5 
)umoles-g wet wlT 1 . 

After 2.5 min of stimulation, pyruvate levels in the muscle increased dramatically 
(Fig. 3). Strombine was not a major glycolytic end product, as this compound accu- 
mulated to levels approaching only 1 /zmole-g wet wt" 1 (Fig. 3). In contrast, octopine 
accumulated linearly during the experiment (Fig. 3, Table II). The sum of octopine, 
free arginine, and arginine phosphate levels was constant (Table II), indicating no 
net change in the total arginine pool in the tissue. Alanopine showed the highest 
accumulation of all the opines, but formation did not begin immediately as this com- 
pound was not detectable in the first 2.5 min of stimulation (Fig. 3). D-lactate was 
the predominant end product formed in the muscle, exhibiting a dramatic increase 
during the later periods of contraction (Fig. 3). 

Octopine, alanopine, and strombine were not released into the medium by the 
muscle preparations. In contrast, a significant amount of D-lactate was found in the 



TABLE II 

Total arginine pool (arginine phosphate, arginine. octopine. and total arginine) for neutralized perchloric- 
acid extracts ofisotonically contracting radula protractor muscles isolated from Busycon contrarium 





Metabolite 


Time 










(min) 


Free arginine 


Arginine phosphate 


Octopine 


Total 





2.37 2.41 


10.69 3.77 


0.46 .29 


13.47 4.39 


2.5 


4.44 1.17 


7.35 2.83 


1.11 .85 


12.55 + 3.34 


5 


3.75 1.04 


6.98 1.50 


1.71 .38 


12.44 1.18 


10 


2.71 .83 


4.93 2. 75 


3.47 1.64 


10.76 2.84 


15 


2.44 .48 


6.98 2.64 


4.72+ 1.73 


14.14 1.26 



Levels are expressed in /umoles - g wet wt ' . Data represent means 



1 SD, n = 5. 



282 



R. W. WISEMAN AND W. R. ELLINGTON 



12. 
11. 

10. 
9. 
8. 
7. 



en .. 
o> 5. 
o 

I 



3. 
2. 

1 . 




ALA 








I 

2.5 




ASP 



10 



15 



2.5 



10 



15 



mm 



FIGURE 2. Changes in levels of the amino acids alanine (ALA), glycine (GLY), and aspartate (ASP) 
as determined by HPLC of neutralized perchloric-acid extracts of electrically stimulated radula protractor 
muscles from Busvcon contrarium over time. Units are ^moles-g wet wt '. Points represent means 1 
SD, n = 5. 



medium (Fig. 4). In fact, during the last 5 min, more D-lactate was released into the 
medium than accumulated in the muscle (Fig. 4). 

DISCUSSION 

The high activities of all four pyruvate reductases in the radula protractor muscle 
of B. contrarium are similar to those observed in the radula retractor muscles of this 
species (Ellington, 1982) and in other gastropod muscles (Baldwin and England, 
1982; Livingstone et ai, 1983). The highest activity, exhibited by octopine dehy- 
drogenase, was comparable to activities observed in cephalopod molluscs (Baldwin 
and England, 1980). The simultaneous accumulation of D-lactate, alanopine, octo- 
pine, and to a lesser extent strombine, which occurs in the radula protractor system 
during muscular activity, indicates that all four of these reductases operate under 
functional anoxia. The accumulation of these end products is temporally correlated 
with changes in pyruvate levels. Pyruvate levels increased dramatically during the 
time course of muscle contraction. 

Pyruvate reductases are thought to be equilibrium enzymes (de Zwaan and 
Dando, 1984; Ga'de and Grieshaber, 1986) and are thus regulated by changes in the 
concentrations of substrates and products. Pyruvate is the common substrate for all 
four reductases. The observed elevations in pyruvate levels in B. contrarium radula 



ANAEROBIC METABOLISM OF RADULA MUSCLES 



283 




mm 



FIGURE 3. Pyruvate (PYR), D-lactate (D-LAC), octopine (OCT, alanopine (ALN), and strombine 
(STR) levels in electrically stimulated radula protractor muscles from Busycon contrarium over time, as 
determined by HPLC and fluorometry. Units are ^moles- g wet wt '. Points represent means 1 SD, n 

c 



protractor would undoubtedly enhance all pyruvate reductase activities. In the case 
of opine dehydrogenases, the concentrations of amino acid co-substrates are also im- 
portant. In the radula protractor muscle, we found that alanine, glycine, and free 
arginine levels were in the 2-8 /umole-g~' range. Finally, pyruvate reductases are 
influenced by accumulation of their respective products. Opine dehydrogenases seem 
to be particularly sensitive to product inhibition (Gade and Grieshaber, 1986). 

D-Lactate was the dominant glycolytic end product even though the maximal 
LDH activity measured in crude tissue extracts was the lowest. Isolated radula muscle 
of B. contrarium displayed the highest levels of D-lactate accumulation yet observed 
during functional anoxia in molluscs. Meinardus and Gade (1981) observed a rela- 



284 



R. W. WISEMAN AND W. R. ELLINGTON 




mm 

FIGURE 4. Distribution of lactate in the incubation media (M) and tissue (Tis) as well as the total 
(To) lactate produced during electrically induced isotonic contractions of the radula protractor muscles 
from the whelk Busycon contrarium as determined by fluorometric analysis of neutralized perchloric-acid 
extracts of each over time. U nits are /imole-g wet wt '. Points represent means 1 SD, n = 5. Where error 
bars are absent, the error bar was less than the size of the symbol used to mark points. 



lively modest accumulation of D-lactate in electrically stimulated preparations of the 
foot muscle of the cockle Cardium edule. The preferential production of D-lactate 
versus the opines in B. contrarium radula protractor muscle may be related to a higher 
binding capacity for pyruvate. The apparent Kms for pyruvate of molluscan LDHs 
are considerably lower than corresponding Kms of ODHs and somewhat lower than 
pyruvate Kms for strombine and alanopine dehydrogenases (Gade and Griesha- 
ber, 1986). 

Isolated radula muscle preparations of B. contrarium did not release octopine, 
alanopine, or strombine into the medium. In addition, the total arginine pool (free 
arginine, arginine phosphate, and octopine) remained constant during contractile 
activity. The decline in aspartate levels during anoxia probably reflects transdeamina- 
tion to alanine because, at the end of 1 5 min, the alanopine-alanine pool size was 
roughly equivalent to the decrease in aspartate plus initial alanine levels. This conser- 
vation of nitrogen in both the arginine (arginine, arginine phosphate, octopine) and 
alanine (alanine, alanopine, aspartate difference) pools is consistent with the observed 
absence of transport of opine end products out of the muscle. 

In contrast to that of the opines, release of D-lactate from the muscle was very 
large. In fact, D-lactate export was greater than D-lactate accumulation during the 
10-15-min period of observation. D-lactate release could have at least two major 
functional advantages. First, removal of the product of this reaction would change 
the mass action ratio ([lactate]/[pyruvate]) in favor of more product formation. Sec- 
ond, if lactate were transported out in a symport system with a proton (H + ) or in an 
antiport system (OH"), this process would help the muscle cells regulate intracellular 
pH 



ANAEROBIC METABOLISM OF RADULA MUSCLES 285 

The total D-lactate produced during the 1 5 min of contractile activity approached 
50 Mmoles-g wet wt '. On the basis of established proton stoichiometries ofglycolysis 
(Portner el al, 1984) i.e., one mole of protons (H + ) per mole lactate (or opine) 
produced lactate and also opine production clearly impose a significant acid load 
on the muscle. Buffering capacities of whelk radular and ventricular muscle, as deter- 
mined by the homogenate titration method (Castellini and Somero, 1981), ranged 
from 30.7 to 39.5 Slykes-g wet wt" 1 (Eberlee and Storey, 1985; Graham and Elling- 
ton, 1985). However, the buffering capacity of B. contrarium ventricles, as deter- 
mined by imposing an acid load (Ellington, 1985) and measuring pH, by phosphorus 
nuclear magnetic resonance ( 3I P-NMR) spectroscopy, yielded a value approaching 
24 Slykes-g wet wt ' (Ellington, unpub. obs.). Regardless of the exact position of the 
buffering capacity, the acid load imposed on the muscle clearly could not be offset by 
purely passive means such as buffering. 

Lactate transport has been studied extensively in erythrocytes. Three mechanisms 
of lactate transport have been identified: (1) non-ionic diffusion, (2) classical anion 
transport, and (3) monocarboxylate carrier (H + symport or OhT antiport) mediated 
transport. Deuticke el al. (1982) reported at least three parallel pathways of lactate 
transport in erythrocytes. Lactate transport in erythrocytes has become the paradigm 
on which mitochondrial (Palmieri el al., 1971), hepatocyte(Fafournoux ?/#/., 1985), 
and whole-muscle (Mainwood and Worsley-Brown, 1975; Seo, 1984) lactate trans- 
port have been modeled. The efflux of lactate from the radula protractor muscle of 
B. contrarium is probably caused by one or several of these mechanisms. Because the 
pKa of lactic acid is 3.86, and the intracellular pH of molluscan tissues under a variety 
of conditions ranges from 7.1 to 6. 6 (Ellington, 1983a; Graham and Ellington, 1985), 
almost all the acid would dissociate into anions; therefore, the rates of non-ionic 
diffusion should be low. A lactate: H + symport (or lactate:OH antiport), if present in 
the radula protractor muscle of B. contrarium, could be of critical importance in 
regulating pHj, especially during periods of elevated glycolytic rates. 

In contrast to lactate, the opines are not released into the medium by the radula 
protractor muscle of B. contrarium. In fact, there is no direct evidence for release of 
opines from any molluscan tissue, although the increase in hemolymph levels during 
contractile activity in cephalopods (Storey and Storey, 1979; Gade, 1980) and the 
decline in the total arginine pool in the mantle muscle during swimming in S. offici- 
nalis (Storey and Storey, 1979) and Loligo vulgaris (Grieshaber and Gade, 1976) 
suggest indirectly that octopine is released in these cases. No other study of contractile 
activity in molluscs has revealed a decline in the total arginine pool or release of 
opines from the muscle (de Zwaan and Dando, 1984). 

Why is lactate readily exported from molluscan muscle cells while opines appear 
to be retained? Specific transporters for amino acids are present in a wide range of 
cell types (Preston and Stevens, 1 982); thus there is no fundamental impediment with 
respect to transport of these compounds. Opine formation results in no net increase 
in the number of osmotically active particles because the amino acid condenses with 
pyruvate derived from a large polymer (glycogen). In contrast, lactate formation re- 
sults in an increase in osmotically active particles, as glycogen is broken down into 
smaller fragments. The lack of disturbance of internal osmolarity has been used as a 
potential functional explanation for the use of opine dehydrogenases rather than 
LDH in certain molluscan muscles (Zandee el al., 1980; Fields, 1983). A logical deriv- 
ative of the argument is that it would be disadvantageous to transport out opine end 
products. However, accumulated end products represent only a small fraction (<5%) 
of the pool of osmotically active substances. Thus, removal of end products, or lack 
thereof, for the sole purpose of cell volume regulation seems unlikely. We favor the 



286 R. W. WISEMAN AND W. R. ELLINGTON 

possibility that the lack of export of opines is related to the lack of a mechanism for 
coupling this movement with regulation of pH,. In fact, since octopine, alanopine, 
and strombine have both positive and negative charges at prevailing pHj conditions, 
a transport mechanism that could couple H + (symport) or OFT (antiport) movement 
with opine export would be difficult to envision. 

The absence of significant amounts of succinate or malate accumulation makes 
the fermentation of aspartate during functional anoxia unlikely. Although the carbon 
skeleton may be unaccounted for, nitrogen is balanced through alanine formation. 
Most probably, the amino group of aspartate is transaminated ultimately to pyruvate 
to yield alanine, with the carbon skeleton of aspartate entering the Krebs cycle as 
malate or oxaloacetate. This hypothesis entails the assumption that there is enough 
oxygen available to the cells to sustain a significant level of aerobic metabolism, at 
least during the early phases of contractile activity. The myoglobin content of radula 
muscle is high (Ball and Meyerhof, 1940; Fange and Mattisson, 1958), and molluscan 
myoglobins typically have low P 50 values (Read, 1 966). Glycolytic rates, as evidenced 
by end product accumulation, are low in the early phases of contractile activity, sug- 
gesting that energy production is largely aerobic during this period. Presumably, the 
oxygen used in this period could be derived from an internal oxygen store such as 
myoglobin. Contractile force decreases in the later portions of the time course while 
pyruvate concentrations increase, suggesting that there is a transition from aerobic 
to anaerobic processes. Thus, aspartate may be an aerobic substrate during the early 
phases of muscle contraction. 

To sum up, our studies have shown that during contractile activity in the radula 
protractor muscle of B. contrarium, high glycolytic rates prevail with pyruvate being 
shunted through all the major pyruvate reductases. Strombine, octopine, alanopine, 
and D-lactate accumulated in order of increasing levels. D-lactate was the major end 
product although D-lactate dehydrogenase displayed the lowest in vitro activity of all 
pyruvate reductases. D-Lactate, but none of the opines, was released from the muscle 
into the incubation medium, with rates exceeding 3 ^moles-min" 1 -g wet wt~'. The 
removal of lactate from the muscle enhances the mass action ratio in favor of lactate 
formation, thereby increasing carbon flux through this enzyme. The removal of lac- 
tate from muscle cells during contractile activity may also help to regulate pH ; 
through a H + symport or OH~ antiport system. This potential role of lactate transport 
in the regulation of pH ; is currently the subject of intensive investigation in this labo- 
ratory. 



LITERATURE CITED 

BALDWIN, J., AND W. R. ENGLAND. 1 980. A comparison of an anaerobic energy metabolism in mantle and 

tentacle muscles of the blue-ringed octopus Hapalochlaena maculosa during swimming. Anst. J. 

Zoo/. 28:407-412. 
BALDWIN, J., AND W. R. ENGLAND. 1982. The properties and functions of alanopine dehydrogenase and 

octopine dehydrogenase from the pedal retractor muscle of Strombidae (Class Gastropoda). Pac. 

Sci. 36:381-394. 
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(Gastropoda:Nassariidae). Mar. Biol. 62: 235-238. 
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CASTELLINI, M. A., AND G. N. SOMERO. 1981. Buffering capacity of vertebrate muscle: correlations with 

potentials for anaerobic function./ Comp. Physiol. 143: 191-198. 
DEUTICKE, B., E. BEYER, AND B. FORST. 1982. Discrimination of three parallel pathways of lactate trans- 



ANAEROBIC METABOLISM OF RADULA MUSCLES 287 

port in human erythrocyte membrane by inhibitors and kinetic properties. Biochim. Biophvs. 

Act a 684: 96- 1 10. 
EBERLEE, J. C, AND K. B. STOREY. 1985. Buffering capacities of the tissues of marine molluscs. Phvsiol. 

Zool. 57, 567-572. 
ELLINGTON, W. R. 1982. Metabolism of the pyruvate branchpoint in the radula retractor muscle of the 

whelk, Busycon contrarium. Can. J. Zool. 60: 2973-2977. 
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luscan tissues: effect of anoxia and ischemia on the intracellular pH and high energy phosphates 

in the ventricle of the whelk, Busycon contrarium. J. Comp. Physiol. 153: 1 59-166. 
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431-444. 
ELLINGTON, W. R. 1985. Metabolic impact of experimental reductions of intracellular pH in molluscan 

cardiac muscle. Mol. Physiol. 7: 155-164. 
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FIELDS, J. H. A. 1983. Alternatives to lactic acid: possible advantages. J. Exp. Zool. 228: 445-457. 
FlORE, G. B., C. V. NICCHITTA, AND W. R. ELLINGTON. 1984. High performance liquid chromatographic 

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139:413-417. 
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aerobiosis. J. Exp. Zool. 228: 415-429. 

GADE, G., AND M. K. GRIESHABER. Pyruvate reductases catalyze formation of lactate and opines in anaer- 
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GRIESHABER, M., AND G. GADE. 1976. The biological role of octopine in the squid Loligo viilgarus (La- 
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Time dependent changes of metabolites in the foot and gill tissue induced by anoxia and electrical 

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K. M. Wilbur and C. M. Yonge, eds. Academic Press, New York. Vol. II. 
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288 R. W. WISEMAN AND W. R. ELLINGTON 

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occurrence of strombine in different organs of the sea mussel Mytilus edulus. Mol. Phvsiol. 2: 

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of the valve snap and valve closure responses in the giant scallop, Placopecten magellanicus. II. 

Biochemistry./ Comp. Physiol. 137: 105-1 15. 



PHYSIOLOGY 

DEATON, LEWIS E. 

Epithelial water permeability in the euryhaline mussel Geukensia 
demissa: decrease in response to hypoosmotic media and hormonal 
modulation +. . . .,.;;. 230 

ENGEL, DAVID W., AND MARIUS BROUWER 

Metal regulation and molting in the blue crab, Callinectes sapidus: met- 
allothionein function in metal metabolism 239 

FELBECK, HORST, AND SANDRA WILEY 

Free D-amino acids in the tissues of marine bivalves .yY. '. 252 

HAND, STEVEN C. 

Trophosome ultrastructure and the characterization of isolated bacte- 
riocytes from invertebrate-sulfur bacteria symbioses . ;"':% .- . /T^A 260 

WISEMAN, ROBERT W., AND W. Ross ELLINGTON 

Energetics of contractile activity in isolated radula protractor muscles 
of the whelk Busycon contrarium: anaerobic end product accumulation 
and release . 277 



,.- '?.- .: 



CONTENTS - ;? 

i 

' ' * 

Annual Report of the Marine Biplp^jpal Laboratory . . , .-. . .-.1 .............. 1 



INVITED REVIEW 

STANLEY-SAMUELSON, DAVID W. 

Physiological roles of prostaglandins and other eicosanoids in inverte- 
brates ____ ........................... . ....... . . . ........ 92 

BEHAVIOR 

CHADWICK, NANETTE E. 

Interspecific aggressive behavior of the corallimorpharian Corynactis 
californica (Cnidaria: Anthozoa): effects on sympatric corals and sea 
anemones ..................... -* ........... . ......... .... 1 1 

DEVELOPMENT AND REPRODUCTION 

BOSCH, ISIDRO, KATHERINE A. BEAUCHAMP, M. ELIZABETH STEELE, AND 

JOHN S. PEARSE 

Development, metamorphosis, and seasonal abundance of embryos and 
larvae of the antarctic sea urchin Sterechinus neumayeri ........... 126 

ECOLOGY AND EVOLUTION 

ALEXANDER, STEPHEN P., AND TED E. DELACA 

Feeding adaptations of the foraminiferan Cibicides refulgens living epi- 
zoically and parasitically on the antarctic scallop Adamussium colbecki 1 36 

BORRERO, FRANCISCO J. 

Tidal height and gametogenesis: reproductive variation among popula- 
tions of Geukensia demissa ................................. 1 60 

MARCUS, NANCY H. 

Differences in the duration of egg diapause of Labidocera aestiva (Co- 
pepoda: Calanoida) from the Woods Hole, Massachusetts, region .... 1 69 

GENERAL BIOLOGY 

HOSE, Jo ELLEN, GARY G. MARTIN, VAN ANH NGUYEN, JOHN LUCAS, AND 

TEDD ROSENSTEIN 

Cytochemical features of shrimp hemocytes .................... 178 

MACKIE, G. O., AND C. L. SINGLA 

Impulse propagation and contraction in the tunic of a compound 
ascidian ............................................... 188 

MANGUM, C. P., K. I. MILLER, J. L. SCOTT, K. E. VAN HOLDE, AND M. P. 

MORSE 

Bivalve hemocyanin: structural, functional, and phylogenetic rela- 
tionships ....... , ....................................... 205 

OKAMURA, BETH 

Particle size and flow velocity induce an inferred switch in bryozoan 
suspension-feeding behavior .......... . ..................... 222 

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IV 



Reference: Biol. Bull. 173: 289-298. (October, 1987) 



ORIENTATION OF THE HORSESHOE CRAB, LIMULUS POLYPHEMUS, 

ON A SANDY BEACH 

MARK L. BOTTOM' AND ROBERT E. LOVELAND 2 

^Fordham University, Divisional' Science and Mathematics, College at Lincoln Center, New York, New 

York 10023, and 2 Rutgers The State University, Department of Biological Sciences and Bureau of 

Biological Research, Nelson Biological Laboratories, Piscataway, New Jersev 08854 

ABSTRACT 

Adult horseshoe crabs (Limulus polyphemus) spawn on sandy intertidal beaches 
and then return toward the water. Field experiments demonstrated that beach slope 
was more significant than vision in this orientation behavior. Both blinded and nor- 
mally sighted crabs showed rapid seaward orientation on beaches with a seaward 
slope of approximately 6. Orientation performance was poor on a flat beach, al- 
though sighted crabs slightly out-performed blinded crabs. The observed orientation 
behavior was correlated with the large numbers of horseshoe crabs which failed to 
return to the water after spawning on sand bars or similar habitats lacking a slope 
gradient. 

INTRODUCTION 

Adult horseshoe crabs (Limulus polyphemus L.) migrate every spring into Dela- 
ware Bay and other Atlantic coast estuaries to spawn on sandy beaches (Shuster, 
1982; Shuster and Botton, 1985). These sublittoral animals find, and amplex with, a 
mate, migrate to the intertidal zone to deposit and fertilize the eggs, and then return 
to the sea. This implies the existence of spatial orientation mechanisms at each critical 
stage of the reproductive cycle. Although little is known about the mechanism of long 
distance migrations from the continental shelf to estuarine spawning grounds (Botton 
and Ropes, 1987), Rudloe and Herrnkind (1976) showed that submerged crabs near 
breeding beaches can orient in response to wave surge. Barlow et al. (1982) found 
that visual cues are important during mating, and Barlow et al. (1986) suggest that 
light may be a significant environmental factor associated with seasonal and diurnal 
variability in mating activity. Visual stimuli may elicit other behavioral responses 
including direction and speed of locomotion (Cole, 1923; Northrup and Loeb, 1923; 
Ireland and Barlow, 1 978) and telson and gill movements (Powers and Barlow, 1 985). 

The orientation cues which enable horseshoe crabs to return to the water after 
mating have not been previously considered. This behavior has important ecological 
consequences because animals "stranded" on an exposed intertidal beach are sub- 
jected to physiologically stressful conditions, including high temperatures, desicca- 
tion, and osmotic imbalance (Herrnkind, 1 983). Among shore zone arthropods, both 
visual cues (e.g., sun, moon, polarized light, landmark orientation) and nonvisual 
cues (such as beach slope) are known (Herrnkind, 1972, 1983;Schone, 1984). In 1985 
during a preliminary beach survey of Delaware Bay in the area of Fortescue, New 
Jersey, we observed large numbers of live horseshoe crabs burrowed on a relatively 
flat section of exposed intertidal beach at low tide. We hypothesized that this aberrant 

Received 27 April 1987; accepted 22 July 1987. 

289 



290 M. L. BOTTON AND R. E. LOVELAND 

behavior resulted from the inability of the animals to orient seaward in the absence 
of beach slope: in this report, we present an experimental test of that hypothesis. 
Orientationa! behavior of sighted and blinded individuals on sloped and flat beaches 
were quantified to evaluate the importance of beach slope and light for adult horse- 
shoe crabs. 

MATERIALS AND METHODS 

All experiments were conducted between 26 May and 7 June 1986 at the New 
Jersey Oyster Research Laboratory (NJORL), located on the Delaware Bayshore in 
Cape May County, New Jersey. The study area was located just north of the labora- 
tory. One site had a slope and sediment composition typical of this relatively uniform 
and undisturbed beach (Shuster and Botton, 1 985). During spring, these beaches have 
a slope of some 6.4 degrees in approximately a westerly direction. The second site 
was a flat sand bar formed by the outlet of a salt marsh creek. It was located approxi- 
mately 100 m north of the sloped beach site, also with the bay toward the west. Horse- 
shoe crabs spawned in large numbers in this and adjacent areas during full moon high 
tides. Both sites within the study area provided similar visual fields: an open horizon 
in the seaward direction and a line of vegetation (mainly Phragmites) above the high 
water line in the landward (easterly) direction. 

Orientation was studied within "arenas" modeled after Mrosovsky and Shettle- 
worth (1968). On each beach, a 4-m radius circle was inscribed and a 30-cm trench 
dug along its perimeter. Stranded horseshoe crabs and any large pieces of debris were 
removed from the arena before use. The arena was divided into 16 equal sectors of 
22.5 each. Sector 1, beginning at compass point north, was arbitrarily designated as 
0, sector 2 began 22.5 in a clockwise direction, et seq. 

Adult male horseshoe crabs were collected from the bay immediately before the 
study of their behavior. Crabs with missing appendages, or with damaged, missing, 
or heavily encrusted eyes, were discarded. Those crabs kept out of water for more 
than 3 minutes were thoroughly wetted down before the trial. The animals used in 
the blinding experiments were prepared by drying the carapace around the lateral 
and median eyes using a paper towel, and then placing patches of opaque adhesive 
duct tape over the eyes. 

Each of the four combinations of beach slope and vision, i.e., sloped beach/ 
sighted, sloped beach/blinded, flat beach/sighted and flat beach/blinded, were run on 
at least two separate days, with a total sample size of not less than 4 1 individuals per 
combination (range, 8-23 crabs per run). Crabs were tested individually to avoid 
behavioral interactions because they often pause or change direction when other 
crabs are encountered along the beach (pers. obs.). To begin a trial, the top few cm 
of sand in the arena was smoothed using a wooden board to remove the track of 
the previous animal; this procedure also disrupted any possible gradients in surface 
sediment moisture. The crab was placed in the center of the arena facing away from 
the water and on a line perpendicular to the shoreline. If the animal burrowed in 
place, it was lifted out of the sand and re-started; if it burrowed twice in succession, it 
was rejected. 

A trial was completed when the animal's prosoma crossed the perimeter of the 
arena. Animals failing to complete a trial, or which burrowed, or which remained 
motionless on the surface of the sediment for 5 consecutive minutes were recorded 
but not used in the statistical analysis of this behavioral data set. Investigators re- 
corded the elapsed time of the test, the section of the arena from which the crab 
exited, and the number of pauses longer than 30 s. The linear distance of the path 
was measured by placing a metered string along the animal's track; a sketch of the 
path for each trial was made. 



ORIENTATION OF INTERTIDAL LIMULUS 291 

TABLE I 

Comparison of the number of "disoriented" (live, rightside-up) Limulus polyphemus on flat and sloped 
beaches in Delaware Bay on each of three days during Spring 1986 

Mean no. disoriented crabs 
per 1 5-m transect 



Date Flat Sloped n 



23 May 


55.2 


12.2 


5 


4.552** 


24 May 


93.3 


9.8 


4 


3.342* 


25 May 


101.5 


4.5 


4 


3.623* 



n = number of replicate 15-m transects counted on each type of beach, t = t-statistic comparing mean 
number of crabs on each type of beach. Means on flat beach significantly greater than mean number on 
sloped beach at P < .05 (*) or P < .005 (**). 



The performance of a group of crabs was assessed using a number of variables 
including percent crossing the perimeter, time required to leave the arena, linear dis- 
tance travelled, and the number of pauses and circles made by the animals. We calcu- 
lated each animal's "meandering score" as the linear distance of the path divided by 
the radial distance (4.0 m) (Mrosovsky and Kingsmill, 1985). We computed mean 
vectors, mean angles of orientation, and 95% confidence intervals for each group of 
animals (Zar, 1984, p. 428). All animals were assumed to cross at the midpoint angle 
in the appropriate sector. The Rayleigh test (Zar, 1984, p. 443) examined the null 
hypothesis that there was no mean population direction. Differences between mean 
angles were analyzed using the Watson-Williams procedure (Zar, 1984, p. 446). 

We surveyed the "disoriented" crabs in transects on the flat and sloped beaches 
on 23-25 May. Disoriented crabs were denned as those live animals remaining right- 
side-up on the breeding beach at low tide, as distinguished from the normal behavior, 
which is to follow the receding tide and spend the low tide period on the intertidal 
sand flat. Live upside-down crabs, "stranded" by wave action, were not counted. 

RESULTS 

Surveys of disoriented crabs 

The number of disoriented (live, rightside-up) stranded crabs was significantly 
higher on the flat beach than the sloped beach on each of the three days (Table I). 
Since similar, or perhaps slightly lower numbers of mating horseshoe crabs actually 
approached the flat beach compared with nearby sloped beaches (based on Botton, 
1982 and personal observations during the 1986 field season), the accumulation of 
disoriented crabs on the flat beach is not likely to be a numerical artifact. Similar 
dense concentrations of disoriented crabs were observed near creek mouths along 
Delaware Bay north of our study area during a survey in 1 985. In contrast, only about 
20-25% of live crabs stranded on sloped beaches during May, 1986 were rightside- 
up (disoriented) animals; the remainder were stranded upside-down as a result of 
wave action and/or telson abnormalities (R. E. Loveland and M. L. Botton, unpub. 
data). Approximately 24-34% of the disoriented individuals on the flat beach were 
stranded as mated pairs (males amplexed to females). By comparison, only 46 disori- 
ented mated pairs (of a total of 4247 stranded crabs) were found on two sloped 90-m 
study beaches between 15 May and 21 June 1986 (R. E. Loveland and M. L. Botton, 
unpub. data). 



292 



M. L. BOTTON AND R. E. LOVELAND 



N 




N 




FIGURE 1 . Orientation behavior of horseshoe crabs with normal vision on a sloped beach. Run A 
held 26 May, 1610-1850, bright sunlight, wind W less than 5 mph. Run B held 2 June, 1057-1323, cloudy 
with occasional drizzle, wind NNE 15 mph. Length and direction of mean vector r shown by solid arrow; 
Rayleigh's Test indicated significant mean population direction in both runs. Seaward direction indicated 
by open arrow. Typical "fish hook" path followed by a crab is shown in A. 



Orientation on sloped beach. All sighted crabs showed strong seaward orientation 
(Fig. 1). The path followed by nearly all crabs in these trials was the "fish hook" 
pattern illustrated in Figure la. Typically, a crab first walked in the direction it was 
placed (in this case, uphill) before turning to its left or right. More crabs turned left 
(n = 26) than right (n = 16) but the difference was non-significant (x 2 - 2.38, .25 

< P < .10). There was no significant difference in mean angle between a group of 
crabs run in late afternoon (n = 23) and a second group (n = 23) run in midday 
(Watson-Williams test, F = 0.229, n.s.). In both trials, 22 animals (96%) completed 
the test and there was no significant difference in the time it took to complete the test 
(Mann-Whitney U-test, z = 0.493, n.s.). The meandering score was significantly 
lower in the late day group (U = 365.0, P < .005). Pausing and circling behaviors 
were noted only five times each (Table II). 

Blinded crabs also showed strong seaward orientation on the sloped beach (Fig. 
2). Mean angles of crabs tested in early morning and mid-afternoon were virtually 
identical. In both the morning and afternoon experiments, 20 animals (87%) com- 
pleted each trial. Animals in the morning trial took significantly longer (Mann- Whit- 
ney U-test, U = 352.5, P < .001) and had a higher meandering score (U = 316, P 

< .002) than the afternoon group. This difference was probably related to a steepening 
of the beach slope before the afternoon trial, which was caused by strong wave action 
several tidal cycles earlier. Blinded animals followed the typical fish hook path de- 
scribed above; the direction of turning was random (20 to the left, 21 to the right). 
No blinded animal circled in either trial and pausing was infrequent (Table II). 

Orientation on flat beach. Horseshoe crabs with normal vision had difficulty ori- 



ORIENTATION OF INTERTIDAL LIMULUS 



293 



TABLE II 

Measures of orientation performance (means and standard errors = SE) of horseshoe crabs 
on sloped and flat beaches 







Sloped 


beach 


Flat beach 


Normal vision 


Blinded 


Normal vision 


Blinded 




Variable 


A 


B 


A 


B 


A 


B 


C 


A 


B 


C 


Time to completion 


88.9 


103.3 


134 


,7 


58.1 


271.9 


128.8 


192.1 


183.2 


395.7 


374.9 


SE 


11.8 


16.4 


19 


.1 


5.6 


53.5 


37.6 


22.0 


61.3 


137.2 


64.8 


Meandering score 


1.21 


1.43 


1, 


.48 


1.37 


2.36 


2.15 


2.01 


1.72 


2.04 


3.45 


SE 


0.03 


1.13 


0.04 


0.07 


0.49 


0.32 


0.15 


0.27 


0.28 


0.37 


No. of pauses 


0.04 


0.26 





22 


0.17 


0.50 


0.50 


0.13 


0.75 


1.87 


1.06 


SE 


0.04 


0.12 





.09 


0.10 


0.20 


0.31 


0.07 


0.41 


0.62 


0.36 


No. of circles 


0.09 


0.13 





.00 


0.00 


1.00 


0.60 


1.17 


0.75 


0.67 


2.33 


SE 


0.06 


0.07 





.00 


0.00 


0.35 


0.31 


0.31 


0.31 


0.21 


0.52 


Number completing 


">T 


22 


20 




20 


11 


8 


21 


6 


6 


13 


Number burrowing 








1 




1 


4 


1 





1 


4 


2 


Number stopped 
























without burrowing 


1 


1 


2 




2 


1 


1 


-> 


1 


5 


3 


Total number in run 


23 


23 


23 




23 


16 


10 


23 


8 


15 


18 



Time to completion (in seconds) and meandering scores based on only those animals crossing the 
perimeter of the 4-m radius testing arena. Letters at top of column designate individual runs. 



enting on the flat beach. Three separate runs were conducted. In afternoon runs un- 
der both overcast and sunlight conditions, values of Rayleigh's z indicated that crabs 
were not significantly oriented in any direction (Fig. 3a, b). In the third run, on a 
sunlit morning, there was significant orientation in a seaward direction (Fig. 3c). 
However, four animals which exited through a seaward-facing sector were actually 
travelling parallel to the shoreline when they crossed the perimeter. 

Sighted crabs on the flat beach had a 10-fold increase in pauses and number of 
circles, and higher meandering scores, compared with animals on the sloped beach 
(Table II). Nine of the 40 animals (22.5%) tested in the three runs either burrowed or 
stopped moving for 5 minutes, thus failing to complete the test. 

Horseshoe crabs behaved differently on the flat beach than on the sloped beach, 
although variability was high among those tested. Very few paths formed the fish 
hook pattern described earlier. Initially, crabs moved rapidly in the direction they 
were placed, but typically they turned and slowed down within the first meter. Many 
crabs reared up on their pedipalps and moved slightly from side to side. This behavior 
was often followed by circling, which in most cases began within 1 m of the release 
point, although some animals made wider loops, "figure 8's," or both (Fig. 3a). 

Blinded crabs were even more disoriented on the flat beach than were the sighted 
individuals. The percentage of crabs completing the test was the lowest of the four 
experimental combinations (Table II). Seven of the 41 crabs (17%) burrowed before 
crossing the perimeter and another 9 (22%) stopped moving without burrowing. Of 
those crossing the perimeter, the direction was random (Fig. 4). There was no signifi- 
cant difference in the time to complete a trial between the three groups; however, 
crabs in the third run had a significantly higher meandering score than those in the 
first two. 

In comparing sighted and blinded crabs on _the flat beach, the blinded animals 
took longer (x = 333.8 s) than sighted animals (x = 201.4 s) to complete the test (U 
= 663, P < .05). The meandering scores were not significantly different between the 



294 



M. L. BOTTON AND R. E. LOVELAND 




N 




N 



7 



7 




FIGURE 2. Orientation behavior of blinded horseshoe crabs on a sloped beach. Run A held 28 May, 
0710-1038, heavy cloud cover with mist, wind WSW 5 mph. Run B held 4 June, 1250-1600, bright sun, 
wind S 10 mph; slightly steeper slope than Run A because of strong wave action during previous 48 hours. 
Length and direction of mean vector r shown by solid arrow; Rayleigh's Test indicated significant mean 
population direction in both runs. 



sighted crabs and the first two groups of blinded crabs, but both were significantly 
lower than the third group of blinded crabs (Kruskal-Wallis test, H = 11.54, P 
< .005). 

DISCUSSION 

As noted by Herrnkind (1983), orientation by shore zone arthropods could poten- 
tially involve visual and/or nonvisual guideposts. Among arthropods, visual cues are 
important in various amphipods, isopods, decapods, insects, and wolf spiders (Herrn- 
kind, 1 972). In contrast, beach slope has been demonstrated to be involved in orienta- 
tion far less frequently (Hamner et ai, 1 968; Craig, 1 973). At our Delaware Bay study 
area, there appears to be a strong visual contrast between the dark horizon in a land- 
ward direction and the open horizon in the bayward direction; therefore, we consid- 
ered the possibility that horseshoe crabs on sandy beaches might be employing vision. 

There is considerable precedent in the literature concerning the behavioral re- 
sponses of Limulus to light. Cole (1923) found that asymmetrically blinded animals 
20-60 mm in diameter (which he erroneously considered to be adults) circled most 
frequently in the direction of the remaining lateral eye. Northrup and Loeb (1923) 
found that young crabs (ca. 1 5 cm length) were negatively phototropic. Limulus can 
also detect polarized light (Waterman, 1950), but at present there is no experimental 
evidence implicating this in any known behavioral response. More recently, Barlow 
and collaborators (e.g., Barlow et ai, 1980) have shown the presence of a circadian 



ORIENTATION OF INTERTIDAL L1MULUS 



295 




N 



N 




N 



/ 




FIGURE 3. Orientation behavior of horseshoe crabs with normal vision on a flat beach. Run A held 
21 May, 1555-1930, heavy cloud cover, wind calm. Run B held 28 May, 1622-1747, bright sun, wind 
WSW 5 mph. Run C held 5 June, 0825-1200, bright sun, wind S 5 mph. Length and direction of mean 
vector r shown by solid arrow; Rayleigh's Test rejected the null hypothesis of no mean population direction 
in Run C only. Seaward direction indicated by open arrow. Typical path followed by a crab is shown in A. 



296 



M. L. BOTTON AND R. E. LOVELAND 



N 




FIGURE 4. Orientation behavior of blinded horseshoe crabs on a flat beach. Runs A (30 May, 1314- 
1450, wind W 5 mph) and B (31 May, 0748-1205, wind W 5 mph) were combined (open circles) because 
of the small number of animals completing each trial. Run C (7 June, 0730- 1 245, mostly sunny, wind S 5 
mph) shown with filled circles; length and direction of mean vector r (solid arrow) based on Run C data. 
Population showed random orientation. Seaward direction indicated by open arrow. 



rhythm in ommatidial morphology and visual sensitivity, modulated by a clock in 
the animal's brain. Visual sensitivity is higher at night, enabling males to recognize 
female "models" even during the evening (Barlow et ai, 1 982). Blinded crabs released 
in the subtidal environment were more disoriented than sighted crabs (Ireland and 
Barlow, 1978). However, both blinded and sighted submerged crabs were capable of 
orienting in the vicinity of the breeding beach when wave surge was present (Rudloe 
and Herrnkind, 1976). 

Experiments with blinded and sighted crabs on both sloped and flat beaches indi- 
cate that beach slope, not visual stimuli, is the primary cue used by horseshoe crabs 
to return to the water. Both blinded and sighted animals showed rapid seaward orien- 
tation on a sloped beach. In contrast, orientational performance was severely im- 
paired on a spawning beach lacking slope. Sighted and blinded crabs showed pausing 
and circling behaviors on the flat beach far more frequently than on the sloped beach. 
Circling, accompanied frequently by the animal's rearing up on their pedipalps, may 
have been an attempt to obtain directional information from gravitational cues. Since 
these directional cues were lacking in the flat arena, it is not surprising that meander- 
ing scores on the flat beach were higher, and many animals either burrowed, stopped 
moving entirely, or left the arena on a heading that took them away from the water. 

The physiological basis for the observed response to beach slope is not clear. De- 
spite the extensive use of Limulus in neurophysiological research, no statocyst or 
other balance organ has been described. However, proprioreceptors have been identi- 
fied from the gnathobases and joints of the walking legs (Barber, 1956, 1960; Pringle, 
1956; Barber and Hayes, 1964). We hypothesize that stimulation of these receptors 
in a crab walking "uphill" may elicit a turning response which then directs crabs 
down the slope, toward the water. A crab can presumably detect when it has started 



ORIENTATION OF INTERTIDAL LIMULUS 297 

"downhill," since once on this heading, turns are infrequent. The information regis- 
tered by these proprioreceptors would be constant on a flat beach, which may under- 
lie their lack of orientation. It seems unlikely that mechanoreceptors on the dorsal 
carapace (Kaplan el al, 1976) or lateral spines (Eagles, 1973) are involved in beach 
orientation, although they may provide positional information under different cir- 
cumstances. 

The comparison between the responses of blinded and sighted animals on the flat 
beach suggests a possible secondary role for vision in sandy beach orientation. One 
of three groups of sighted crabs showed significant seaward orientation (Fig. 3), but 
why this group did, and the other two did not, is not apparent. Overall, sighted crabs 
on the flat beach exhibited somewhat better orientation (as measured by percent fin- 
ishing, number of pauses, and meandering scores) than the blinded crabs on the same 
arena, although their performance was much poorer than the blinded crabs on the 
sloped beach. 

A comparable and ecologically similar orientation problem confronts female sea 
turtles after egg laying. In these animals, in contrast to Limulus, the primary cues are 
visual. Females, as well as hatchlings, assess the brightness differential between the 
open, seaward horizon and the darker, vegetation-lined landward horizon (Ehrenfeld 
andCarr, 1967; Mrosovsky and Carr, 1967; Mrosovsky and Shettleworth, 1968;Mro- 
sovsky, 1970). In comparison, there is some evidence for positive geotropism among 
hatchlings (Parker, 1922; Van Rhijn, 1979), although Mrosovsky and Kingsmill 
(1985) point out that on many turtle nesting areas, slope is very irregular and not as 
good a predictor of seaward direction as is the open horizon. This configuration is 
markedly different at the Cape May shore of Delaware Bay, where undisturbed 
beaches consistently have a seaward slope. A positive geotaxis is therefore a reliable 
orientation behavior for adult horseshoe crabs, whereas for sea turtles, it is not. How- 
ever, it might be adaptive for a species to have a secondary orientation mechanism 
should the primary system fail. For sea turtles, geotaxis may function, albeit weakly, 
when vision is impaired (Van Rhijn, 1979); a similar hierarchy of cues exists in the 
amphipod, Orchestoidea corniculata (Herrnkind, 1983). For horseshoe crabs, vision 
may be of some limited value when the crabs become stranded on a flat beach, but 
many animals which spawned on this type of beach are unable to find the water 
only a few meters away (Table I). Under these circumstances, it seems adaptive for 
horseshoe crabs to burrow because it conserves energy and keeps their book-gills in 
contact with cooler, moist sand. If high tide during the next day is of equal or greater 
amplitude than the one on which they were stranded, the crabs will have a good 
chance of survival, but if tidal heights are declining, there may be localized, large- 
scale mortality. 

ACKNOWLEDGMENTS 

We are grateful to Marine Biologicals, Inc., and the Fordham University Research 
Council for financial support, and to Dr. R. A. Lutz for the use of laboratory facilities. 
We are also grateful to K. A. Becker, P. Claxon, and P. Jones for their assistance in 
running the behavioral trials. 

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BARBER, S. B. 1960. Structure and properties of Limulus articular proprioreceptors. J. Exp. Zool. 143: 

283-321. 
BARBER, S. B., AND W. F. HAYES. 1964. A tendon receptor organ in Limulus. Cornp. Biochem. Physiol. 

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298 M. L. BOTTON AND R. E. LOVELAND 

BARLOW, R. B., JR. S. C. CHAMBERLAIN, AND J. Z. LEVINSON. 1980. Limulus brain modulates the struc- 
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BARLOW, R. B., JR., L. C. IRELAND, AND L. KASS. 1982. Vision has a role in Limulus mating behaviour. 

Nature 296: 65-66. 
BARLOW, R. B., M. K. POWERS, H. HOWARD, AND L. KASS. 1986. Migration of Limulus for mating: 

relation to lunar phase, tide height, and sunlight. Biol. Bull. 171: 3 10-329. 
BOTTON, M. L. 1982. Predation by adult horseshoe crabs, Limulus polyphemus (L.) and its effect on ben- 

thic intertidal community structure of breeding beaches in Delaware Bay, New Jersey. Ph.D. 

thesis, Rutgers University. 466 pp. 
BOTTON, M. L., AND J. W. ROPES. 1987. Populations of horseshoe crabs on the northwestern Atlantic 

continental shelf. Fish. Bull, (in press). 

COLE, W. H. 1923. Circus movements of Limulus and the tropism theory. /. Gen. Physiol. 5: 4 1 7-426. 
CRAIG, P. C. 1973. Orientation of the sand-beach amphipod, Orchestoidea corniculata. Anim. Behav. 21: 

699-706. 
EAGLES, D. A. 1973. Lateral spine mechanoreceptors in Limulus polyphemus. Comp. Biochem. Phvsiol. 

44A: 557-575. 
EHRENFELD, D. W., AND A. CARR. 1967. The role of vision in the sea-finding orientation of the green 

turtle (Chelonia rnydas). Anim. Behav. 15: 25-36. 
HAMNER, W. M., M. SMYTH, AND E. D. MULFORD JR. 1968. Orientation of the sand-beach isopod Tylos 

punctatus. Anim. Behav. 16: 405-409. 
HERRNKIND, W. F. 1972. Orientation in shore-living arthropods, especially the sand fiddler crab. Pp. 1- 

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Press, New York. 
HERRNKJND, W. F. 1983. Movement patterns and orientation. Pp. 41-105 in The Biology of Crustacea, 

Vol. 7, Behavior and Ecology. F. J. Vernberg and W. B. Vernberg, eds. Academic Press, New 

York. 
IRELAND, L. C., AND R. B. BARLOW JR. 1978. Tracking normal and blindfolded Limulus in the ocean by 

means of acoustic telemetry. Biol. Bull. 155: 445-446. 
KAPLAN, E., R. B. BARLOW JR., S. C. CHAMBERLAIN, AND D. J. STELZNER. 1976. Mechanoreceptors on 

the dorsal carapace of Limulus. Brain Res. 109: 6 1 5-622. 
MROSOVSKY, N. 1 970. The influence of the sun's position and elevated cues on the orientation of hatchling 

sea turtles. Anim. Behav. 18: 648-65 1 . 
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turtles, Chelonia mydas. tested on their natural nesting beaches. Behaviour 28: 2 1 7-23 1 . 
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water finding behavior of sea turtles. Behaviour 27: 21 1-257. 
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PARKER, G. H. 1922. The crawling of young loggerhead turtles toward the sea. J. Exp. Zool. 36: 323-33 1 . 
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PRINGLE, J. W. S. 1956. Proprioreception in Limulus. J. Exp. Biol. 33: 658-667. 
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SCHONE, H. 1984. Spatial Orientation. Princeton University Press, Princeton, New Jersey. 347 pp. 
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Reference: Biol. Bull. 173: 299-310. (October, 1987) 



THE FEEDING BEHAVIOR OF PAR.4NOPHRYS CARNIVORA 
(CILIATA, PHILASTERIDAE) 

DAVID KAHAN. 1 THEODORA BAR-EL. 1 NORBERT WILBERT, 3 
SAMSON LEIKEHMACHER, 2 AND SAMUEL OMAN 2 

1 Department of Zoology, Hebrew University of Jerusalem, Jerusalem, Israel, 2 Department of Statistics, 

Hebrew University oj Jerusalem, Jerusalem, Israel, and *Zoologisches Institut, 

Poppelsdorfer Schloss, Bonn, West Germany 

ABSTRACT 

The marine ciliate Paranophrys carnivora, isolated from the Mediterranean coast 
of Israel, was found to feed on a varied diet of bacteria, algae, and living and non- 
living tissues. Chlorococcum sp. and Duualiella parva, the algal species on which P . 
carnivora grew best, did not elicit a chemosensory response; tissues and bacteria did. 
In experiments on stationary phase ciliates, betaine, choline, L-histidine, and tri- 
methylamine oxide elicited a positive chemosensory response at concentrations as 
low as 10~ 6 Mto 10" 3 A/. 

INTRODUCTION 

Most ciliates feed on particulate matter consisting mainly of microorganisms 
(bacteria and algae) of a size appropriate to their buccal apparatus. The particles, 
whether suspended or settled, are collected via specialized cilia near the oral opening 
(Corliss, 1979). 

In Tetrahymena, one of the ciliates most studied, particulate matter seems obliga- 
tory for feeding. An autoclaved 2% proteose peptone medium which supports a 
flourishing culture, loses this capability when the particles are removed by millipore 
filtration. Addition of inert particles lacking any nutritive value to the filtered me- 
dium restores its growth potential (Rasmussen and Kludt, 1970; Rasmussen and 
Modeweg-Hansen, 1973). Extensive experiments performed by Fenchel ( 1980a, b, c) 
with inert "latex" particles indicate that various ciliates select their food primarily by 
particle size. 

When offered different combinations of algal species of the same size range, the 
ciliate Favella ehrenbergii showed a preference for one species, indicating that food 
selection is also based on factors other than size (Stoecker el al., 1981). Selective 
feeding has been attributed to chemical stimuli in various other ciliates e.g., Nassula 
(Poilvert, 1959), Stentor coeruleus (Tartar, 1961; Rapport et al., 1972), and Podo- 
phrya calkinsi (Hull, 1 96 1 ). In addition, particle movement affects the feeding behav- 
ior of ciliates (Karpenko et al., 1977). 

Most studies on the feeding behavior of ciliates focused on those feeding on micro- 
organisms e.g., Parameeiiim, Tetrahymena (Levandowsky and Hauser, 1978; Van 
Houten et al., 1981, 1982; Antipa et al., 1983; Levandowsky et al., 1984; Leick and 
Hellung-Larsen, 1985; Hellung-Larsen et al., 1986). Studies on ciliates that feed on 
tissues are scarce (Levandowsky and Hauser, 1978; Van Houten et al., 1981). The 
marine ciliate Paranophrys carnivora, which was recently described, feeds on tissues 
of living or dead organisms (Czapik and Wilbert, 1986). The feeding behavior of this 
organism is described in this paper. 

Received 9 February 1987; accepted 22 July 1987. 

299 



300 D. KAHAN ET AL. 

MATERIALS AND METHODS 
Cultivation and morphology of Paranophrys carnivora 

P. carnivora was isolated from samples collected from the Mediterranean Sea at 
the coast of Dor, Israel. The samples were rich in various protozoans: Ciliata e.g., 
Euplotes, and Flagellata, mostly autotrophic ones, including Dunaliella. Initial obser- 
vations showed that P. carnivora fed on algae. They were also observed to gather in 
the vicinity of freshly injured invertebrates (crustaceans) and feed on their tissues. 

Several P. carnivora clones were prepared and grown on Dunaliella parva as well 
as on a strain of the bacterium Enterobacter aerogenes which can grow at a salinity 
of 35%. No attempt was made to eliminate the original bacterial flora. The most suc- 
cessful clone was further cultured on E. aerogenes for growth curve and feeding be- 
havior studies. These bacteria were grown on brain heart Agar (Difco) slants at 28C, 
and then harvested in sterile water to give a suspension having an absorbance of 1 . 1- 
1.3 at 430 nm, as measured with a Bausch and Lomb "Spektronic 20" spectropho- 
tometer. A 2-ml inoculum of a 3-4 day-old culture of Paranophrys and 0.4 ml of the 
bacterial suspension were added to a test tube (3 cm diameter, 20 cm length) contain- 
ing 30 ml of sterile (autoclaved) 35%o artificial seawater (Instant Ocean salts from 
Aquarium Systems, Mentor, Ohio, in filtered water, hereafter referred to as ASW). 
The culture was then incubated at 28C in a temperature-controlled chamber. For 
growth curve studies, 0.5- or 1-ml samples were removed at various intervals from 
each of three cultures which had been inoculated simultaneously from the same 
source, and preserved in a 10% Bouin solution. All of the ciliates in each sample were 
counted in a glass chamber at 60X with a hand tally. Size determinations were made 
at 600X using an ocular micrometer. 

The morphological description and identification was based on the same clone 
which was kept in 15 ml of ASW in covered glass vials at 20C and fed every 14 
days on 3 pin-head sized bits of either oligochaete or crustacean meat. Biometric 
measurements were made using a light microscope on ciliates stained by the protargol 
(Wilbert, 1975) and silver nitrate (Chatton and LwofF, 1930) methods. For scanning 
electron microscopy, cells were fixed instantaneously by rapid addition of a large 
volume of 2.7%. OsO 4 in ASW. After 10-15 minutes in fixative at room temperature, 
the cells were washed with 2% glutaraldehyde in ASW. After 10 minutes they were 
dehydrated in a graded ethanol series, dried by the critical point method, coated with 
gold-palladium, and viewed in a Joel 840 scanning electron microscope. 

Feeding and growth experiments with marine algae 

The species of marine algae whose names are given in Table II were cultivated in 
test tubes in a medium of ASW enriched with Walne solution (Walne, 1966) in a 
temperature-controlled room at 18C and under continuous illumination. Young, 
flourishing week-old cultures were inoculated with Paranophrys carnivora and fur- 
ther incubated under continuous illumination at 25C. The cultures were examined 
with a dissecting microscope at 40X both initially and at various intervals during a 
week to appraise the ciliate population growth. Samples were also examined under 
higher power of the light microscope, while alive and after they were killed with a 1% 
formalin solution. At 60X, the dimensions of the algae (length/width or diameter) 
were determined using an ocular micrometer. Ciliates were also examined for vacu- 
oles containing algae. 

Behavior experiments 

Capillary tube assay. The amino acids tested were purchased from Sigma (L- 
leucine, L-isoleucine, L-proline, hydroxy-L-proline, L-arginine, D- and L-histidine, 



FEEDING BEHAVIOR OF P. CARNIl'OR.4 301 

L-cysteine); Nutritional Biochemical Corporation (DL-phenylalanine, DL-a-alanine. 
DL-serine, L-methionine, L-threonine, DL-asparagine); British Drug House, Ltd. 
(glycine, L-aspartic acid); Merck & Co., Inc. (L-tyrosine); Light & Co., Ltd. (L-cys- 
tine); Fluka (DL-valine); and CHR (L-tryptophan). Betaine hydrochloride, choline 
chloride, and trimethylamine oxide (TMAO) were also purchased from Sigma, and 
proteose peptone and brain heart infusion from Difco Laboratories. 

The substances to be tested were dissolved in distilled water and the pH of the 
solution was adjusted, if necessary, to the pH of ASW (pH 8). Glass capillary tubes 
(Modulohm of Helver, Denmark) 5-8 mm long and 0.7-1.0 mm in external diame- 
ter, were filled with the test solution and dried. In a previous study with another 
species of the same family, Porpostoma not at um, many ciliates would enter a control 
capillary containing fresh medium even when the medium they were swimming in 
was only half-an-hour old (Kahn et ai, 1981). Therefore we modified the capil- 
lary assay by drying the test solutions and then, during the experiment, allowing the 
test substances to become dissolved in the same medium the ciliates were swimming 
in, rather than use fresh medium as a solute. Control capillary tubes were not filled 
with any chemicals. Tissue culture dishes, 35 X 10 mm (Falcon), were each filled with 
2 ml of ciliates from the stationary phase of culture, which had been diluted with fresh 
ASW to a density ranging from 100 to 300 ciliates per ml. For each concentration and 
substance to be tested, a test and a control tube were placed in different halves of each 
experimental dish. When both test and control dry capillaries were immersed in the 
ciliate suspension at the start of each experiment, they became filled with the medium 
in which the ciliates were suspended. The test substances were dissolved within the 
experimental period (the substances found later to elicit a positive chemosensory re- 
sponse were further tested separately in a series of identical capillaries. These sub- 
stances dissolved completely within a five-minute period). In each experimental run, 
a test and control capillary pair were tested in each of three dishes i.e., in triplicate. 
Betaine, found earlier to elicit a chemosensory response from Paranophrys carnivora 
(mistakenly identified as P. magna: Kahan et ai, 1985), was used at a concentration 
of 10~' M (together with a control capillary) as a standard in each experimental run 
to verify the responsiveness of the ciliates. These betaine and control pairs were also 
run in triplicate. Using a dissecting microscope, the number of ciliates in each tube 
(up to about 100) was recorded with a hand tally at intervals during a 30-minute 
period. Levandowsky et al. (1984) preferred using flat capillaries to eliminate diffi- 
culties encountered in viewing Tetrahymena ciliates through cylindrical capillaries. 
We did not experience difficulty in counting moving Paranophrys carnivora in the 
cylindrical tubes. In the initial screening, most of the amino acids were tested at a 
concentration of 10 ' M, with the exception of L-glutaminc acid and L-tryptophan, 
at 5 X 10" 2 M, and L-tyrosine, at 2 X 10~ 3 M. L-histidine was the only amino 
acid which elicited a positive response at least as strong as that of betaine. This amino 
acid, as well as betaine, choline, and trimethylamine oxide (TMAO), was further 
tested in at least four experimental runs at concentrations from 10~ 6 or 10~' 
to 10" 'A/. 

The chemosensory response was computed at each time interval as the ratio of 
the number of ciliates in the tube containing the test substance to the total number 
of ciliates in both the test and control tubes. Since statistical analysis (via Mests) of 
the differences in response at different times during each half-hour experimental run 
showed no consistent effect of observation times, the index of chemosensory response 
for a given experimental run was defined as the maximum of the chemosensory re- 
sponses at the time intervals measured. To adjust for variation in chemotactic respon- 
siveness over the different days of the experimental runs, a relative index of chemotac- 
tic activity for a given substance at a given concentration, was defined as the ratio of 



302 D. KAHAN ET AL. 

its chemotactic activity to the index of the standard (betaine, at a concentration of 
10" 1 M) for the same experimental run. 

Both the index and relative index of chemosensory response of P. carnivora, for 
the various substances at different concentrations, were analyzed by two-way analysis 
of variance (Scheffe, 1959). Effects on the index due to substances or concentrations 
were analyzed using the S-method of multiple comparisons (Scheffe, 1959). 

Dialysis experiments. Dialysis bags, 20 cm in length and 1 .6 cm in diameter ( Visk- 
ing Tubing, The Scientific Instrument Center, Ltd.), were filled with 10 ml of either 
test solution (5% proteose peptone in ASW) or control (ASW alone). They were im- 
mersed in separate finger bowls each containing 150 ml of ciliate suspension at a 
density of 40 per ml. This was prepared by diluting a stationary phase culture with 
fresh ASW. To evaluate the behavioral effect, the tubing was first examined along 
its entire length using the low magnification of the dissecting microscope for the 
greatest congregation of ciliates. This section was further examined under 40X and 
the number of ciliates on both the test and control bags was compared at consecutive 
time intervals for up to 2 hours. 

RESULTS 

The growth curve of Paranophrys carnivora and associated morphological changes 

The growth curve of P. carnivora fed on Enterobacter aerogenes at 28C is shown 
in Figure 1. Figure 1 shows that the logarithmic growth phase continues for up to 
about 30 hours with a generation time of 7-8 hours. The stationary phase which 
follows is short, and after 48 hours there is a moderate decline in the number of 
ciliates. This phase continues until the experiments are terminated at the end of the 
fifth day. During the growth experiments the shape of the cell changed from ovoid 
(the "trophic" form, having a length to width ratio of 1.8 in the logarithmic phase) 
to more elongated (the "swimming" form, with a ratio of 2.2 or more in the stationary 
and decline phases). More pronounced differences between the two forms were ob- 
tained with cultures fed on either oligochaete or crustacean meat and, rarely, from 
cultures fed on algae. Scanning electron micrographs of the two forms from cultures 
fed on Dunaliella parva are shown in Figures 2 and 3. The biometric data presented 
in Table I are of silver stained specimens from cultures fed on oligochaete or crusta- 
cean meat, as are the light micrographs given in Figures 4 and 5. In Figure 5, note 
the marked appearance of the stained kinetosomes and the protrichocysts, another 
characteristic of the swimming form. 

Feeding and growth experiments with marine algae 

Table II shows that Paranophrys carnivora ingested most of the algae offered. 
However, different results were obtained with the various algal species ingested. The 
best growth was obtained with Chlorococcum sp. and Dunaliella parva; no growth 
occurred with Chlorella saccharophila and Dunaliella primolecta. As might be ex- 
pected, those algal species that were not ingested did not support good ciliate cultures. 

Chemosensory response 

In the laboratory, P. carnivora fed on either Enterobacter aerogenes, various algae, 
or wounded Anemia, dead or alive, when each of these diets was offered individually. 
Differences in chemosensory responses were obtained when capillary tubes contain- 
ing one of the diets was offered with a capillary containing no food (control), and the 
number of ciliates in each of the two tubes compared after 10 minutes. As shown in 



FEEDING BEHAVIOR OF P. CARNIl'OR.4 



303 



10 



ou 
O 



o> 

CL 



CD 



0.5 



o 

i_ 

CL> 
_Q 

E 

13 




20 40 60 80 100 120 
Hours 

FIGURE 1 . Growth curve of Paranophrys carnivora. Results are based on three replicate cultures. 



Table III, Artemia homogenate and the E. aerogenes suspension elicited a positive 
chemosensory response, whereas the alga Dunaliella parva elicited none. These re- 
sults were obtained with ciliates that had been previously cultivated on each of the 
diets indicated. When Chlorococcum sp. was offered instead of D. parva, the same 
results were obtained. 

Since Enterobacter was routinely cultivated on brain heart agar, it was thought 
that its effect could have been due to the presence of some dissolved ingredients from 
the growth medium in the suspension. Indeed, brain heart infusion did elicit a posi- 
tive chemosensory response in capillary experiments. To determine whether the bac- 
teria themselves are effective, they were washed by centrifugation and offered to the 
ciliates in a capillary tube. After repeated rinsings in ASW, neither the bacterial pellet 
nor the supernatant gave a positive result. However, when the washed pellet was 
suspended in fresh ASW, incubated for up to 48 hours at 1 8C, and then centrifuged 
again, the resulting supernatant elicited a chemosensory response. This indicates that 
washed bacteria excrete with time an effective substance or release such a factor upon 
disintegration. 

Positive results were obtained with other microbiological growth media, such as 
proteose peptone and casein hydrolysate. In dialysis experiments with proteose pep- 
tone, Paranophrys was found to be attracted to those molecules which were able to 
pass through the cellophane membrane, e.g., amino acids. In further capillary tests 
to screen individual amino acids, only L-histidine, of the various amino acids tested, 
was as effective as betaine, a substance previously found to elicit a chemosensory 
response from P. carnivora (Kahan et al, 1985). Choline and trimethylamine oxide 
(TM AO), compounds with a chemical structure similar to that of betaine, also elic- 
ited a positive response at least as strong as that of betaine. The four substances (beta- 



304 



D. KAHAN ET AL. 









FIGURES 2-5 are to the same scale and view the ventral side (note buccal cavity). 

FIGURE 2. Scanning electron micrograph of trophic form ofParanophrys carnivora (arrowhead indi- 
cates part of an algal cell, Dunaliella parva, engulfed by the ciliate). 

FIGURE 3. Scanning electron micrograph of swimming form of P. carnivora (arrowheads indicate 
algal cells from culture which have adhered to the ciliate). 

FIGURE 4. Photomicrograph of silver-stained trophic form of P. carnivora. 

FIGURE 5. Photomicrograph of silver-stained swimming form of P. carnivora. 



FEEDING BEHAVIOR OF P. CARNIVORA 305 

TABLE I 

Cell dimensions of the trophic and swimming forms of Paranophrys carnivora (given in micrometers) 



Length 


Width 




Distance from 
anterior pole 














to end of UM 


Form 


Range 


x S.E. 


Range 


.vS.E. 


Range 


;cS.E. 


Trophic 


36-56 


47.08 1.3 


18-35 


25.18 1.07 


18-25 


20.82 1.17 




(17) 




(22) 




(17) 




Swimming 


40-60 


49.44+ 1.47 


13-22 


16. 67 0.62 


24-29 


26.4 + 0.52 




(18) 




(15) 




(15) 





Numbers in parenthesis indicate the number of observations. 



ine, L-histidine, choline, and TM AO) were offered to P. carnivora at various concen- 
trations; the results were analyzed as described previously. The index averaged over 
the experimental runs is shown in Figure 6 for each substance at the various concen- 
trations. The average index of chemosensory response elicited by betaine, choline, 
and TMAO was significantly greater than 0.5, at the 5% level or more, at all the 
concentrations examined, and by L-histidine for concentrations of at least 10~ 3 M 
(concentrations of 10~ 4 , 10 5 , and 10" 6 M were also tested, but the response was not 
significantly greater than 0.5). 

Two-way analysis of variance of the index of chemosensory response showed sig- 
nificant effects due to material (P-value : : 0.028) and concentration (f-value : 
0.000). The means of the relative index of chemosensory response (the index, pre- 
viously defined under Materials and Methods as "capillary tube assay," which adjusts 
for the level of response to the betaine standard during the same experimental run) 
exhibited the same behavior as the nonadjusted index, indicating that the differences 
in the index of chemosensory response represented in Figure 6 are not due to varying 
levels of overall chemosensory responsiveness (as measured by the response to the 
standard, betaine at 10~' A/) on the different days of experimental runs. The analysis 
of variance is based on the nonadjusted chemosensory response, as opposed to the 
adjusted response, because the data on the former satisfied the required statistical 
assumptions on the error terms more closely (see Scheffe, 1959, p. 5). 

Figure 6 suggests that choline, betaine, and TMAO are similar in the response 
they elicit from P. carnivora, the main differences in the chemosensory response in- 
dex being due to different concentrations. This was confirmed for betaine and choline 
using the S-method (Scheffe, 1959) of multiple comparisons to compare effects on 
the chemosensory response index due to materials or concentrations. The average 
index for betaine and choline at the concentration of 10~' M was significantly differ- 
ent (at the 5% level) from the average at the 10~ 6 M level; and the differences between 
the averages at the 10~' M and 10~ 5 M levels, and at the 10 2 M and 10" 6 M levels, 
were almost significant at the 10% level. 

DISCUSSION 

Although previously thought (Czapik and Wilbert, 1986) to feed on fresh and 
decomposing tissues, the present study establishes that Paranophrys carnivora can 
feed on a more varied diet. This diet includes algae and bacteria in addition to tissues. 
In this respect it seems to be closer in its dietary spectrum to P. thompsoni and P. 
magna than to the other species of the genus. P. thompsoni was reported to live on 
bacteria and heterotrophic flagellates which developed in hatched gelatinous egg 



306 



D. KAHAN ET AL. 



TABLE II 
Growth on and ingestion of different algal species by Paranophrys carnivora 



Algal class: 
Species 


Source 


Algal size" 


Ingestion Ciliates' 
of algae growth 6 


Baciilariophyceae 








Amphora sp. 12 


J.L.,CUNY C 


35-40/2-4 


no + 


Phaeodactylum 








tricornutum 


CMBRDG CC d 


20-30/2.5 


yes ++ 


Chlorophyceae 








Chlamydomonas 








provasoli 


J.L.,CUNY 


4-8 


no + 


Chlamydomonas 








hedleyi 


J.L.,CUNY 


5-10/3-9 


yes + + 


Chlorella 








stigmatophora 


IOLR e 


5-6/3-4 


yes + 


Chlorella 








saccharophila 


IOLR 


3 


yes 


Chloroccoccum sp. 


J.L..CUNY 


2-3 


yes ++++ 


Dunaliella 








primolecta 


CMBRDG CC 


6-14/4-13 


yes 


Dunaliella sp. 








Strain C9AA 


B.G.,HU r 


10-18/8-13 


no + 


Dunaliella sp. 








Strain El 


B.G., HU f 


14-18/8-10 


no + 


Dunaliella sp. 








Strain 1644 


B.C., HU f 


11-21/8-15 


no + 


Dunaliella sp. 








Strain L10 


B.C., HU f 


9-14/8-13 


no + 


Dunaliella 








tertiolecta 


B.G.,HU f 


8-12/4-8 


yes ++ + 


Dunaliella parva 


B.G., HU f 


6-12/3-8 


yes ++++ 


Dunaliella sp. 








Strain 14 


E.G., HU f 


6-10/3-8 


yes + + + 


Dunaliella sp. 








Strain E4 


B.C., HU f 


5-10/3-8 


yes + + + 


Dunaliella sp. 








Strain Iran 6 


B.C., HU f 


5-8/3-8 


yes + + + 


Nannochloris sp. 








Strain W5 15 


J.L.,CUNY 


8-12/6-8 


no + + 


Cyanophyceae 








Anacystissp. 


Houde 8 


2-3 


yes + + 


Prasinophyceae 








Tetraselmis chuii 


IOLR 


12-13/8-9 


no 


Prymnesiophyceae 








Isochrysis galbana 


IOLR 


4-6 


yes ++ 



a The dimensions (length/width or diameter) are given in ^m. 

"Rating code: ++++, excellent; +++,good; ++, fair; +, poor; -, no growth. 

c John Lee, City University of New York. 

d Cambridge Culture Collection. 

e Institute of Oceanographic and Limnological Research, Haifa. 

f B. Ginzburg, Hebrew University (Ginzburgand Ginzburg, 1985). 

8 E. D. Houde, University of Miami, Florida. 



masses of dipterans (Didier and Wilbert, 1976), while P. magna was cultivated in 
cultures to which split peas had been added (Borror, 1972) and presumably fed on 
the bacterial flora. Nevertheless, comparative dietary experiments on the above-men- 
tioned species should be further extended in order to establish their feeding pattern. 



FEEDING BEHAVIOR OF P. CARNIVOR.4 



307 



TABLE III 

Chemosensory response of Paranophrys carnivora to different diets as determined by capillary assay 

Test diet offered in capillary 



Diet cultivated on 



Dunaliella 
parva 



Enterobacter 
aerogenes 



Anemia 
homogenate 



Algae (Dunaliella parva) 
Bacteria 

(Enterobacter aerogenes) 
Fresh meat 

(wounded Anemia) 






Cultures of the ciliates were grown each on a different diet as indicated. The "+" indicates a positive 
response and " ", no response. 



Our attempts (unpub.) to introduce P. carnivora as a symbiont living in the coelenter- 
ates Cordylophora sp., Cassiopea sp., and Aiptasia as well as in the crustaceans Ar- 
ternia salina and Macrobrachium rosenbergii, did not succeed. Two other species of 
the genus, P. marina and P. carcini, were found inside coelenterates (Thompson and 



x 

CD 



CD 

CO 

C 

o 

CL 
tO 
CD 



co 

C. 
CD 
CO 
O 



CD 

-C 


C 
D 
CD 



.0 



0.9 



0.8 



0.7 



0.6 



0.5 




10" 



10' 



,-2 



10"- 10 ' 10"- 10 
Molar concentration 



_c 



FIGURE 6. Dose-response curves for substances which elicit a positive chemosensory response from 
Paranophrys carnivora by the capillary assay. Each figure represents the mean of the values of the index 
of chemosensory response obtained from all of the observations for a particular substance at a certain 
concentration. The results obtained with betaine are indicated by circles, with trimethylamine oxide by 
triangles, with choline chloride by squares, and with L-histidine by diamonds. A full (black) figure indicates 
that the mean index of chemosensory response is significantly greater than 0.5 at the 1% level, and a half- 
full figure, at the 5% level. L-histidine was also tested at 10 4 , 10~ 5 , and 10~ 6 M. but the results were not 
significantly greater than 0.5. 



308 D. KAHAN ET AL. 

Berger, 1965) and in the hemolymph of crustaceans (Groliere and Leglise, 1977), 
respectively. 

Like many other ciliates (Fenchel, 1980a, b, c), P. carnivora ingests suspended 
inert particles such as polystyrene beads (unpub.) and living microorganisms. Here 
size seerns to be a limiting factor in food ingestion. The largest food vacuoles observed 
did not exceed 7 yum in diameter, and algal species having size ranges above this 
limit were not ingested i.e., Amphora sp., Dunaliella strains C9AA, El, 1644, L10, 
Nannochloris sp., and Tetraselmis chuii. However, some of the large species (the four 
strains of Dunaliella mentioned above and Nannochloris sp.) did sustain growth. This 
could be due to the ciliates' feeding on bacteria contaminating the algal cultures and/ 
or on disintegrating algal cells in aged cultures. The same explanation could be offered 
for the ciliates' growth on Phaeodactylum triconutum. While C. provasoli was in the 
size range of algae that could be engulfed, it was not ingested. This is probably due to 
the tendency of the latter algal cells to form bigger sized aggregates, or to their having 
a chemoinhibitory effect on phagocytosis by Paranophrys. Those species of algae that 
were ingested by Paranophrys (Table II) gave growth results that varied in their rating 
from no growth, i.e., Chlorella saccharophila and Dunaliella primolecta, to excellent 
growth i.e., Dunaliella parva and Chlorococcum sp. However, these latter two species 
did not elicit a positive chemosensory response from Paranophrys carnivora in our 
experiments. Although algae are known to release assimilated carbon into the culture 
medium (Hunstman, 1972; Fogg, 1977; Saks, 1982), Dunaliella parva and Chlorococ- 
cum sp. evidently do not release a substance eliciting a chemosensory response from 
Paranophrys carnivora. 

Betaine, choline, L-histidine, and trimethylamine oxide, the substances found to 
elicit a positive chemosensory response from Paranophrys carnivora, are known to 
affect feeding behavior in various other organisms (Lindstedt, 1971; Levandowsky 
and Hauser, 1978; Heinen, 1980; Caprio, 1984). They are also widely distributed in 
various organisms including bacteria and algae (Bell and Mitchell, 1972; Levandow- 
sky and Hauser, 1978; Edwards, 1982; Galinski and Truper, 1982; Abe, 1983; Ko- 
nosuetal.. 1983; Shirani et a/., 1983; Imhoffand Rodriguez-Valera, 1984; Morihiko 
et al, 1984) and therefore could be indicators of a wide variety of food sources for 
Paranophrys carnivora. There may be other effective substances as yet untested for 
Paranophrys carnivora, which have recently been found to elicit a chemosensory re- 
sponse from other ciliates such as Parameciwn (Antipa and Norton, 1985) and Tetra- 
hymena (Leick and Hellung-Larsen, 1985; Hellung-Larsen et al., 1986). Paranophrys 
carnivora responds to the D-isomer of histidine, which does not occur in nature, and 
in this respect resembles Tetrahymena thermophila ( Almagor et al., 1 98 1 ), which also 
responds to both the L and D forms of an amino acid (methionine). 

Another characteristic Paranophrys carnivora shares with Tetrahymena is its 
body transformation. The morphological differences in body proportions between 
the ovoid feeding form and elongated swimming form of P. carnivora appear more 
pronounced when the ciliate is cultivated on tissues and on rare occasions when 
grown on algae, after depletion of the food organisms. A similar transformation in 
form appears after starvation in Tetrahymena thermophila. In the latter, the transfor- 
mation is accompanied by several other changes i.e., oral replacement, caudal cilium 
appearance, and increase in number of somatic basal bodies and cilia, as well as in 
speed (Nelsen, 1978; Nelsen and DeBault, 1978). In P. carnivora, significant changes 
in body proportion and an increase in the somatic basal bodies have been noticed. 
Greater control of culture conditions of the ciliates (as may be obtained with an axe- 
nic culture) would enable further discerning and understanding of this phenomenon 
in P. carnivora. 



FEEDING BEHAVIOR OF P. CARNIVOHA 309 

ACKNOWLEDGMENTS 

D. K. gratefully acknowledges the Schonbrunn Foundation for funding the re- 
search performed at the Hebrew University of Jerusalem. The assistance of Eli Hatab, 
M. Devorachek, and A. Nevo in photographing SEM pictures is greatly appreciated. 
We are grateful to M. Levandowsky for reading the manuscript and making valuable 
comments. 

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Reference: Biol. Bull. 173: 31 1-323. (October, 1987) 



INTRACELLULAR pH DECREASES DURING THE IN VITRO 

INDUCTION OF THE ACROSOME REACTION IN 

THE SPERM OF SICYONIA INGENTIS 

FRED J. GRIFFIN, WALLIS H. CLARK JR., JOHN H. CROWE, AND LOIS M. CROWE 

Department of Zoology, University of California, Davis, California 95616 and Bodega Marine 

Laboratory, Bodega Bay, California 94923 

ABSTRACT 

Activation of the sperm of many invertebrate and some vertebrate species to un- 
dergo an acrosome reaction is accompanied by an increase in intracellular pH (pH ; ). 
In each of these instances the pHj of the unactivated cell is relatively low (6.9-7.4). 
Unactivated sperm of the marine shrimp, Sicyonia ingentis, possess an elevated pHj 
(8.5). Induction of the acrosome reaction (exocytosis of the acrosomal vesicle and 
generation of an acrosomal filament) is accompanied by a decrease in pH, (7.8). Low 
external pH elicits acrosomal filament formation in sperm that have undergone acro- 
somal exocytosis, but does not induce exocytosis in unreacted sperm. The ionophore, 
nigericin, enhances the percent of sperm that form filaments in low pH seawater (pH 
< 8.0), but does not elicit filament formation at external pHs > 8.0. Valinomycin 
induces filament formation in sperm that have undergone exocytosis over a wide 
range of external pHs (5.75-8.5). The ability of valinomycin to induce filament for- 
mation in the upper portion of this pH range (8.0) declines as the extracellular K + 
concentration rises. These results demonstrate that the sperm of S. ingentis undergo 
a pHj decrease as a result of the acrosome reaction and that the decrease is associated 
with acrosomal filament formation. In addition, they also suggest that an efflux of K + 
ions is connected to the pHi decrease. 

INTRODUCTION 

As a prerequisite to fertilization, most sperm must first undergo acrosomal alter- 
ations, termed the acrosome reaction (Dan, 1952). Among the majority of inverte- 
brate sperm and sperm of a few select vertebrates, the acrosome reaction (AR) is 
composed of the exocytosis of the acrosomal vesicle and generation of an acrosomal 
filament (reviewed by Dan, 1967; Austin, 1968). The AR occurs when a sperm con- 
tacts an egg-derived inducer. The inducer, a component of one of the egg investments, 
interacts with the plasma membrane of the sperm and initiates a series of ionic and 
biochemical sperm-associated events that lead to, among other sperm-associated 
changes, the AR (see Shapiro and Eddy, 1980; Lopo, 1983 for reviews). For example, 
the ionic changes associated with the AR in the sperm of Strongylocentrotus purpura- 
tus include: ( 1 ) an uptake of extracellular Ca ++ which is thought to be involved in the 
exocytosis of the acrosomal vesicle (Tilney et at, 1978; Shackman et at, 1981); (2) a 
Na + associated H + efflux which is necessary for the polymerization of actin filaments 



Received 29 May 1987; accepted 28 July 1987. 

Abbreviations: ASW (artificial sea water), DMO ( l4 C-dimethyloxazolidine 2,4-dione), EW (egg water), 
pH, (intracellular pH), pH (extracellular pH). 

Reprint requests to: W. H. Clark, P.O. Box 247, Bodega Bay, CA 94923. 

311 



312 F. J. GRIFFIN ET AL. 

and thus, the formation of the acrosomal filament; and (3) a K + efflux that leads to a 
depolarization of the sperm membrane potential (Shackman el ai, 198 1 ). Although 
not thoroughly documented, it appears that the ARs of many motile invertebrate 
sperm involve the same ionic changes. 

Unlike the sperm of most invertebrates, the sperm of the natantian decapod, Sicy- 
onia ingentis, are nonmotile. These cells possess an anterior spike (contained within 
an acrosomal vesicle), a subacrosome, and a posteriorly located main body which 
houses the nucleus (Kleveet al, 1 980; Shigekawa and Clark, 1986). S. ingentis sperm 
do not possess flagella and also lack organized mitochondria. Sperm are transferred 
to female seminal receptacles during mating and stored until spawning, which may 
occur several weeks to months later (Anderson el al., 1985). Thus, these sperm re- 
main in an unactivated state for extended periods after transfer from the male. At 
spawning, ova are released from paired ovopores and mixed with sperm ejected from 
the seminal receptacles. Sperm bind spike first to ova and become activated to un- 
dergo a biphasic AR (Clark el al., 1984). Within seconds bound sperm undergo the 
first phase of the AR, acrosomal exocytosis (which includes the loss of the spike), 
and 10-20 minutes later they complete the AR by generating an acrosomal filament 
(second phase). Thus, in vivo the two phases of the AR are temporally separated. 
Previous work demonstrates that the first phase, acrosomal exocytosis, depends upon 
external Ca ++ (Clark el al., 198 1 ) as is true in other systems. These experiments were 
conducted with sperm taken from males, which are not competent to form acrosomal 
filaments (Clark el al., 1984). The ionic requirements for acrosomal filament forma- 
tion have not been investigated. 

The ability to induce a complete AR in sperm removed from female seminal 
receptacles and incubated in isolated egg products enabled us to investigate the ionic 
requirements of the AR's second phase, acrosomal filament formation. The present 
paper: ( 1 ) describes the in vitro induction of a complete AR using egg components; 
(2) presents data suggesting that the sperm possess a high intracellular pH (pHj) prior 
to undergoing the AR and that a pH, decrease is associated with the second phase of 
the AR (formation of the acrosomal filament); and (3) provides data indicating that 
the outward movement of K + ions is involved in the pHj drop. 

MATERIALS AND METHODS 
Collection and maintenance of animals 

Specimens ofSicyonia ingentis were collected using an otter-trawl in 60-100 me- 
ters of water off San Pedro, California. Live animals were transported in chilled sea- 
water (8-10C) to the Bodega Marine Laboratory and maintained in flow-through 
seawater tanks at ambient temperatures ( 10-14C). Gravid females were isolated and 
kept under constant light in a 500 gallon flow-through tank. The lights were turned off 
to initiate spawning. Animals were monitored for spawning under a red light (Kodak 
Wratten #2 filter). 

Collection of egg water 

Spawning animals were removed from the tank and held over 50 ml glass beakers 
containing chilled (4C) artificial seawater (ASW) prepared according to Cavanaugh 
(1956). After the negatively buoyant ova (1-2.5 X 10 3 ) had settled to the bottom 
of the beaker, approximately 3 /4 of the seawater was drawn off. The ova were then 
resuspended by swirling and kept in suspension for five minutes. The remaining ASW 



A pH, DECREASE AT SPERM ACTIVATION 313 

containing egg-derived components was then pipeted out of the beakers, cleared by 
centrifugation (100,000 X g, 15 min), and divided into 1 ml aliquots. The protein 
concentration of each egg water (EW) batch was determined after the method of 
Lowry et al. (1951). EW was stored in liquid nitrogen if not used immediately. 

Collection of sperm 

In S. ingentis, only sperm that have been transferred to a female and stored in the 
female's seminal receptacles are competent to: (1) undergo the acrosome reaction 
(AR) in response to egg derived components; and (2) form an acrosomal filament as 
part of the AR, regardless of the manner of induction (Clark et al., 1984). As a result, 
only sperm taken from seminal receptacles of the female were used. Seminal recepta- 
cles from ten or more females were pooled, homogenized in ASW using a Wheaton 
5 ml tissue grinder to free sperm, and hand centrifuged to remove fragments of empty 
receptacles. Free sperm were pelleted from the supernatant at 200 X g for five min- 
utes. Pelleted sperm were resuspended in ASW and used within one hour of isolation. 

Induction of the acrosome reaction with egg water 

Isolated sperm ( 10 6 cells) were incubated in 1 ml of experimental (containing EW) 
and control (containing ASW) solutions. Aliquots of cells in each experiment were 
fixed (with a drop of 5% glutaraldehyde in ASW) at appropriate times and scored with 
phase microscopy (400X) for: ( 1 ) percent unreacted; (2) percent that had undergone 
acrosomal exocytosis but had not formed acrosomal filaments; and (3) percent fully 
reacted (sperm which possessed acrosomal filaments). For each experimental run (n), 
duplicate 20 n\ aliquots were removed and 100 sperm were scored in each for acroso- 
mal status. 

Intracellular pH determinations 

Isolated sperm (2. 1 X 10 8 ) were divided into three equal samples. One sample was 
incubated for 10 min in 1.6 ml of ASW (pH 8.0); this sample was used to measure 
the pHj of unreacted sperm. A second sample was incubated in 1 .6 ml of EW (pH 7.8) 
for 10 min; these sperm were used to measure the pHj of sperm that had undergone 
acrosomal exocytosis. The last sample was incubated in 1 .6 ml of EW for 50 min; 
these sperm were used to measure the pHj of fully reacted sperm. Intracellular pH 
determinations were made with 14 C-dimethyloxazolidine 2,4-dione (DMO) using a 
modification of the technique described by Waddell and Butler ( 1 959). At the conclu- 
sion of the initial incubations, each of the three samples was divided in half: ( 1 ) 25 /A 
of 3 H 2 O and 50 iA of 14 C inulin were added to one; and (2) to the other, 25 n\ of 3 H 2 O 
inulin and 50 n\ of 14 C DMO (final concentration of 33 ^M) were added. After a 20 
min equilibration period: (1) triplicate 20 ^1 aliquots (controls) were transferred to 
scintillation vials containing 15 ml of ACS scintillation fluid (Beckman); (2) 10 /A 
samples were removed, fixed, and scored for acrosomal status; and (3) triplicate 200 
/tl samples were microfuged (Fisher Model #235B) for 90 seconds through a 95 vol- 
ume percent silicone oil (Dow Corning 704)-5 volume percent hexane solution, the 
supernatants were removed, and the tips of the microfuge tubes (containing the sperm 
pellets) were cut off and placed in scintillation fluid. The samples sat overnight and 
were then counted on a Beckman LSI 00 scintillation counter. Calculation of internal 
water space and pH { determinations followed those described by Shackman et 
al. (1981). 



314 F. J. GRIFFIN ET AL. 

pH induction experiments 

Sperm (10 7 /ml) were incubated for 5 min in either ASW pH 8.0 or EW pH 8.0 
after which 100 ,ul aliquots were added to 900 ^1 of ASW at the following pHs: 5.75, 
6.0, 6.5, 7.0, 7.5, 8.0, 8.5, 9.0, and 9.5. Reactions were halted 10 min later with a 
drop of 5% glutaraldehyde (in ASW) and acrosomal status was scored. In experiments 
using sperm that had spontaneously undergone acrosomal exocytosis in ASW alone, 
sperm were scored until 100 reacted cells had been observed in each duplicate. In 
those using EW to induce acrosomal exocytosis, counts were performed as described 
above. The pH of ASW was determined on an Orion (EA920) pH meter. Above pH 
8.0, ASW was adjusted with 0.1-1.0 TV NaOH; below pH 8.0 it was adjusted with 
0.1-1.0 A^HCl or 0.2 Macetate buffer. All ASWs were pH adjusted just prior to use. 

lonophore induction experiments 

In separate experiments, sperm (10 7 /rnl) were induced to undergo acrosomal 
exocytosis in EW as described above and 100 n\ aliquots were added to 900 jul of 
ASW at the pHs described in the previous section. Immediately after the addition of 
sperm, 5 ^1 of nigericin (0.5 mA/in 100% DMSO) or valinomycin (1.0 mMin 100% 
DMSO) were added, with mixing, to the sperm suspensions. Control samples con- 
tained 0.5% DMSO. Aliquots of each treatment were fixed with a drop of 5% glutaral- 
dehyde (in ASW) 5 minutes after the addition of the ionophores or DMSO and sperm 
were scored for acrosomal status. In addition, samples were removed and fixed to 
determine levels of acrosomal exocytosis prior to introduction into pH ASWs and 
the addition of the ionophores. 

ASWs of different [K + ] were obtained by adding or deleting equal molar amounts 
of KC1 and NaCl from the MBL formula for ASW (Cavanaugh, 1956). 

RESULTS 

Induction of the acrosome reaction by egg water 

Acrosomal status of S. ingentis sperm is easily scored with phase microscopy. 
Figure 1 illustrates an unreacted sperm, a sperm that has undergone acrosomal 
exocytosis, and a fully reacted sperm (possessing an acrosomal filament). Sperm re- 
moved from the seminal receptacles of females and incubated in 50 ng/m\ (protein) 
of egg water (EW) undergo a complete AR (acrosomal exocytosis and formation of 
an acrosomal filament) in which the temporal separation between the two phases is 
maintained. Within 1 min of exposure to EW, S. ingentis sperm underwent acroso- 
mal exocytosis at levels greater than ASW controls (41% as compared to 7.3%) and 
by 5 min, greater than 75% of the sperm had undergone acrosomal exocytosis (Fig. 
2). These sperm (after a 1 5 min incubation in EW) had only undergone acrosomal 
exocytosis; they did not possess acrosomal filaments. Sperm that had undergone acro- 
somal exocytosis did begin to form filaments, commencing approximately 30 min 
after introduction into EW (Fig. 3). At 45 min, more than 50% of all sperm counted 
possessed acrosomal filaments; this translates into more than two-thirds of exocy- 
tosed sperm possessing filaments. By 60 min, approximately 60% of all sperm (85% 
of exocytosed sperm) had formed filaments. The sperm that formed acrosomal fila- 
ments were those that had undergone acrosomal exocytosis. The total percent of 
sperm that had either undergone only exocytosis or undergone a complete AR re- 
mained constant through 60 min (Fig. 3). 



A pH, DECREASE AT SPERM ACTIVATION 



315 




FIGURE 1. Phase micrographs of the three activational states of Sicyonia ingentis sperm; (A) an 
unreacted sperm possessing an anterior spike, (B) a sperm that has undergone acrosomal exocytosis and 
has lost the spike, and (C) a fully reacted sperm possessing an acrosomal filament. 



A small but consistent number of sperm (8-10%) isolated in artificial seawater 
(ASW), and not transferred to EW, underwent acrosomal exocytosis (Fig. 2). This 
percent not only remained constant with increased incubation times, but exocytosis 
was the only portion of the AR that occurred. Sperm isolated to ASW have been 
observed for up to 1 80 min without seeing acrosomal filaments. 

Intracellular pH measurements 

Based on the accumulation ratios of the DMO uptake experiments we have calcu- 
lated an average pH; for unreacted and reacted S. ingentis sperm. Unreacted sperm 



100, 



80- 



CO 

o 60 

o 
o 
x 40 - 

Ld ^ U ~ 



20- 












-4- 



5 10 

TIME (MIN) 



15 



FIGURE 2. EW induction of acrosomal exocytosis; response over time. Sperm were incubated in 50 
^g/ml EW (O) or ASW (), fixed at the times designated above, and scored for acrosomal status. Data 
points are means; vertical lines are standard deviations (n = 4). Each replicate utilized a separate batch of 
sperm and EW. 



316 



F. J. GRIFFIN ET AL. 



t/2 
in 
o 

i 

o 
o 

X 

LJ 



100- 



80- 



60- 



40- 



20- 







-I 




15 30 

TIME (MIN) 



45 



r 100 



-80 



-60 



-40 



-20 







60 



m 
z 
i 

CO 



FIGURE 3. Acrosomal filament formation as a function of time. Sperm were incubated in 50 
of EW. Aliquots of sperm were removed, fixed, and scored for percent exocytosis (O) and percent formed 
filaments () at each time point. Data points are the means of four replicates; each replicate was conducted 
with different batches of sperm and EW. Vertical lines are standard deviations. 



removed from seminal receptacles and placed in ASW (pH 8.0) possessed a pH ; of 
8.47 0.27. In these samples, greater than 91% of the sperm were unreacted and 
none of the reacted sperm possessed acrosomal filaments (Table I). Sperm incubated 
in EW(pH 7.8) for 10 min prior to the addition of the DMO possessed a significantly 
lower pHj of 7.81 0.13 (P < 0.05). Greater than 74%. of these sperm had undergone 
acrosomal exocytosis and approximately 3% possessed acrosomal filaments at the 
end of the equilibration period (30 min after sperm had been introduced into EW). 
The pH, of sperm incubated in EW for 50 min, 8.01 0.06, was also lower than that 
of unreacted sperm (P < 0.05), but was not significantly different from the 10 min 
EW samples. Seventy-one percent of the sperm incubated in EW for 50 min possessed 
filaments at the end of the equilibration period (70 min after introduction into EW). 



TABLE I 



of Sicyonia ingentis sperm 



Sample 



Exocytosed' 



Filaments 2 



pH, 



ASW 
EW,o 
EW 50 



8.3 2. 5 

72. 3 3.1 

6.0 2.0 





2.7 1.5 
71.0 2.0 



8.47 0.27 
7.81 0.13 
8.01 0.06 



Isolated sperm were reacted in EW for 10 min (EW, ) and 50 min (EW 50 ) and used to measure pH, of 
sperm that had undergone only acrosomal exocytosis and sperm that had fully reacted, respectively. The 
pH, of unreacted sperm (ASW) was measured after incubating isolated sperm for 10 min in ASW. 

' Percent of sperm which had undergone acrosomal exocytosis only at the time of disruption. 

2 Percent of sperm which had undergone a complete AR at the time of disruption. 



A pH, DECREASE AT SPERM ACTIVATION 317 

Effect of external pH on theAR 

External pH (pH ), within the range examined, does not elicit the first phase of 
the AR, acrosomal exocytosis, in S. ingentis sperm (Fig. 4 A). The percentages of 
sperm that were unreacted after transfer to the pH ASWs did not vary significantly 
with pH . This was true for those sperm that had been preincubated in ASW alone, 
as well as for those sperm that had been preincubated in EW. In those experiments 
where sperm had been preincubated in EW, the percent that did not undergo acro- 
somal exocytosis averaged 26. 1 for all pH s with no observable pH-dependent trend. 
The same held true for those sperm that were not exposed to EW (incubated in ASW 
only); the percent of these sperm that did not react averaged 90.7 for all pH treat- 
ments. Such was not the case with regard to acrosomal filament formation. 

Exposure of sperm to low pH did elicit the formation of acrosomal filaments. 
Sperm induced to undergo exocytosis with EW and subsequently transferred to pH s 
of less than 7 underwent filament formation within 10 min of transfer (Fig. 4B). This 
represents a reduction of the temporal separation between the two phases of 20-35 
min. The pH optimum for filament induction was between pH 5.75 and 6.5. Below 
pH 5.75, exocytosed sperm were disrupted and above pH 7.0, sperm did not form 
acrosomal filaments within the reduced temporal window. Similar results were ob- 
tained with sperm that had spontaneously undergone exocytosis in ASW (not incu- 
bated in EW). The percentages of these sperm that formed filaments after exposure 
to low pH were somewhat less than the sperm treated with EW, but the effect of pH 
was similar (Fig. 4). 

Induction of acrosomal filament formation by ionophores 

The response of sperm to low pH was enhanced with the addition of either nigeri- 
cin or valinomycin, however, the pH optima were different for both ionophores (Fig. 
5). Greater than 50% of sperm that had undergone acrosomal exocytosis in EW and 
were subsequently exposed to nigericin for 5 min underwent filament formation in 
pH 6.0-7.5. At pH 5.75, 31.5 8.2% of such sperm possessed filaments, however, 
the number of unreacted sperm was twice that of the other pH treatments, suggesting 
that at pH 5.75 reacted sperm were disrupting (Fig. 5A). With the addition of nigeri- 
cin, not only were the percentages of filament formations increased at low pH s (6.0- 
7.5), but the range for filament induction was shifted 0.5-1.0 units basic. 

Valinomycin not only elicited more filament formations than any of the other 
treatments, but it was also effective over a broader range of pH s than the other treat- 
ments (Fig. 5). Greater than 80% of exocytosed sperm underwent filament forma- 
tion in pH 5.75-8.0. At pH s 9.0 and 9.5 the percentages of filaments were dramati- 
cally reduced. As in low pH inductions in the absence of ionophores, there was no 
observable effect on acrosomal exocytosis (Fig. 5A). 

The ability of the ionophore valinomycin to induce formation of acrosomal fila- 
ments was pH dependent when extracellular K + was elevated (Fig. 6). Filament for- 
mation in pH 6.0 ASW was not reduced by increasing concentrations of extracellular 
K + , however, a steady decline in the percentage of filaments was observed in pH 8.0 
ASW as the K + level was increased. When the [K + ] was increased from 10 mA/to 20 
mM, filament formation at pH 8.0 decreased by approximately 50%. At 30 mM K + 
filament formation declined by another 50%, and at 40 mM K + no filaments were 
observed. In pH 9.0 ASW, filaments were only formed in low K + (K + -free ASW). 



318 



F. J. GRIFFIN ET AL. 



Q 
LJ 

I 
O 

LJ 

rr 

I 

o 



100n 

80- 
60- 
40 
20 







9 







8 



10 



CO 

h- 

LJ 



100n 



80- 



60- 



B 




PH 

FIGURE 4. Effects of external pH upon the AR. (A) Acrosomal exocytosis. (B) Acrosomal filament 
formation. Sperm ( 10 7 /ml) were incubated for 5 min in either ASW pH 8.0 () or EW pH 8.0 (O) after 
which 100 ^1 aliquots were added to 900 ^1 of ASW at the pHs indicated above. Reactions were halted 10 
min later with a drop of 5% glutaraldehyde (in ASW) and acrosomal status was scored. Data points in (B) 
represent mean % of exocytosed sperm that formed filaments. 



DISCUSSION 

Induction of the two phases (acrosomal exocytosis and acrosomal filament forma- 
tion) of the AR in S. ingentis sperm is temporally separated and sequential in vivo 
(Clark et al., 1984). Upon binding to ova, sperm undergo acrosomal exocytosis and 
some 10-20 min later undergo acrosomal filament formation. The present report has 
demonstrated that the in vitro induction of this AR in sperm removed from female 



A pH, DECREASE AT SPERM ACTIVATION 



319 



O 
< 

Ld 

o: 

I 

z 
o 



80- 
60- 
40- 
20- 





8 



10 



CO 

i 

z: 

UJ 



B 




PH 

FIGURE 5. The effects of pH on valinomycin and nigericin induction of the AR. (A) Acrosomal 
exocytosis. (B) Acrosomal filament formation. Sperm (10 7 /ml) were first induced to undergo acrosomal 
exocytosis in EW, 100 n\ aliquots were added to 900 ^1 of ASW at the pHs indicated above, and then 
exposed to either 1 /uA/ nigericin (O) or 5 nM valinomycin (). Sperm were fixed after 5 min and scored for 
reactions. Data points in (B) represent mean percent of exocytosed sperm that formed acrosomal filaments; 
vertical lines are standard deviations, n = 3. Each n in each experimental batch represents sperm pooled 
from different females, different ASWs, and different EW and nigericin solutions. 



seminal receptacles and incubated in solutions containing isolated egg components 
(EW) is also temporally separated and sequential. In vitro, acrosomal exocytosis is 
achieved within 2.5-5 min, yet sperm that have undergone acrosomal exocytosis do 
not form acrosomal filaments for an additional 30-45 min. Thus the temporal sepa- 
ration that is observed on the surface of an ovum is preserved albeit lengthened under 



320 



F. J. GRIFFIN ET AL. 



100-1 



PH 6.0 




20 
K + (mM) 

FIGURE 6. Acrosomal filament formation and external K + . Sperm (10 7 /ml) were induced to un- 
dergo acrosomal exocytosis in EW (5 min). One hundred (100) ^1 samples were then transferred to 900 n\ 
of ASWs containing from to 40 mM K + at pH 6.0 (A), pH 8.0 (), and pH 9.0 (A). Samples were fixed 
at 5 min and scored for acrosomal status. Control samples (O) were preincubated in ASW (not exposed to 
EW), added to ASW (pH 8.0) containing the described [K + ], and exposed to 0.5% DMSO. Data points are 
mean percent of exocytosed sperm that formed filaments (n = 3); vertical lines are standard deviations. 



in vitro conditions. The ability to elicit a complete AR in vitro and the fact that the 
two phases are separated has allowed the dissection of the two phases with respect to 
the controls of activation. Based upon direct measurements of pHj, low pH induc- 
tions of the AR, and the effects of both low pH and external [K + ] on ionophore 
inductions, we propose that formation of the acrosomal filament in 5". ingentis sperm 
is associated with a pH, decrease. 

Measurements of intracellular pH in S. ingentis sperm suggest that: ( 1 ) unreacted 
sperm possess a high intracellular pH; (2) prior to formation of the acrosomal fila- 
ment these cells undergo a pHj decrease; and (3) subsequent to filament formation 
they do not return to the unactivated pH,. Although DMO is a widely used probe for 
determining pHj, it does have limitations (Roos and Boron, 1981; Busa and Nucci- 
telli, 1 984). These include: ( 1 ) DMO measurements reflect an average pH for the cell 
and do not provide information on the pH of subcellular compartments (e.g., the 
acrosomal vesicle or the subacrosome); and (2) alkaline membrane-bound organelles 
can sequester DMO, giving an erroneous picture of the pH of other subcellular com- 
partments (Roos and Boron, 1981; Busa and Nuccitelli, 1984). For example, 
Grinstein el al. (1984) have reported that a DMO measured pHj increase at lympho- 
cyte proliferation is in fact not an activational pHj change, but rather an increase in 
the number of mitochondria (which results in an increased DMO uptake by the cells). 
The structural organization and the direction of the measured pHj change in S. 
ingentis sperm, however, allowed us to entertain the supposition that the pHj change 
was real and was associated with filament formation. Unreacted S. ingentis sperm 
possess three subcellular regions: a nucleus, a subacrosome, and an acrosomal vesicle; 



A pH, DECREASE AT SPERM ACTIVATION 321 

mature sperm do not possess mitochondria (Shigekawa and Clark, 1986). As a result, 
any pH; changes would be expected to be associated with one of these compartments 
and two of them are involved in the AR. We would not expect, a priori, an overall 
pHj decrease to occur simply as a result of acrosomal exocytosis; the acrosomal vesicle 
is an acidic organelle (Kleve et al, 1980) and therefore its loss at exocytosis might be 
expected to yield an increase in average pH, . It was therefore reasonable to expect the 
pHj changes to be associated with the subacrosome. 

Results of the low pH induction experiments correlate well with the pH, measure- 
ments and delineate at which phase of the AR the pH ; drop occurs. Neither low 
pH (<7.5) alone nor low pH in conjunction with nigericin or valinomycin induce 
unreacted sperm to undergo acrosomal exocytosis. All three do induce acrosomal 
filament formation in sperm that have undergone exocytosis. It follows that the pH ; 
decrease is associated with the second phase of the AR, formation of the acrosomal 
filament. Furthermore, low pH elicits filament formation in sperm that have exocy- 
tosed in ASW and have not been exposed to EW. This indicates that the pH is not 
acting through a pH alteration of EW, rather, it is directly influencing filament forma- 
tion. These observations are in contrast to previous studies demonstrating that a net 
rise in pH ; occurs during the AR in sperm of other species (Shackman et al., 1981; 
Working and Meizel, 1983; Matsui et #/.,1986). By contrast with the sperm of S. 
ingentis, these cells have been reported to possess depressed pH,s prior to activation. 
For example, the pH; of unreacted S. piirpuratus sperm is between 6.6 and 7.3, based 
upon measurements obtained with weak bases (Shackman et al., 1981). Using 9- 
aminoacridine, the pH, of the sperm of the starfish Aster ias amurensis and A. pectini- 
fera was reported to be 7.4-7.5 (Matsui et al., 1986). In the hamster sperm, the intra- 
crosomal pH has been measured to <5, also using 9-aminoacridine (Meizel and 
Deamer, 1978). 

The ionophore nigericin exchanges K + or Na + for H + (the selectivity for K + over 
Na + is more than an order of magnitude), thus it is an electroneutral ionophore that 
dissipates proton gradients across cell membranes (Pressman, 1976; Johnson and 
Scarpa, 1976). As such, the pH, of sperm in the presence of nigericin should more 
closely parallel the pH of the ASW than in the low pH experiments conducted 
without ionophore. The results of the nigericin induction experiments agree well with 
the measured pH ; changes that occur during the AR. Based on the DMO measure- 
ments, sperm decrease pHj from 8.5 to between 7.8-8.0 as a result of the AR. Nigeri- 
cin elicits filament formation at pH s 6.0-8.0 in sperm that have undergone 
exocytosis. Since the pH /pHj gradient at pH s above 8.0 would not favor a nigericin 
induced pHj decrease, filament formation would not be expected. Conversely, as the 
pH is decreased, it would be expected that at some pH an acid overload in the 
presence of nigericin would occur. This occurs between pH 5.75 and 6.0 in S. ingen- 
tis sperm. 

Valinomycin, like low pH ASW and nigericin, does not elicit acrosomal ex- 
ocytosis, but will induce acrosomal filament formation. However, unlike the other 
two, valinomycin is pH-independent over a wide pH range (pH 5.75-8.0) at normal 
extracellular K + concentrations (10 mA/). The ability of valinomycin to elicit fila- 
ment formation does become sensitive to pH at elevated extracellular K + concentra- 
tions. In 10 mA/ K + ASW filament formation proceeds at pH 6.0 and 8.0; no fila- 
ments are seen at pH 9.0. As the [K + ] is increased to 40 mM(in 10 mA/ increments), 
filament formation declines approximately 50% at each incremental rise in [K + ] in 
pH 8.0 ASW. At pH 9.0, filament formation is inhibited in [K + ] > 10 mA/, however, 
filament formation will proceed if the K + concentration is below 10 mA/. Valinomy- 



322 F. J. GRIFFIN ET AL. 

tin transports only K + (the selectivity over Na + is greater than three orders of magni- 
tude) across r ::mbranes and therefore is electrogenic (Johnson and Scarpa, 1976; 
Pressman, 1 976). The results of the valinomycin/pH/K + experiments suggest that the 
ionophr re is facilitating a K + efflux, however, they also suggest that the K + efflux 
does not in itself elicit filament formation. The fact that filament formation in pH 
8.0 ASW is very sensitive to small changes in the extracellular K + concentration leads 
us to suggest that the pHj decrease elicits acrosomal filament formation. The iono- 
phore facilitates a K + efflux which results in an alteration of the sperm membrane 
potential (hyperpolarization?) and this change in membrane potential drives a proton 
influx. 

This study has demonstrated that unactivated sperm of S. ingentis possess an 
unusually high resting pHj, that they undergo a decrease in pH, as a result of the AR, 
and that the pHj decrease is associated with formation of the acrosomal filament. The 
pHj measurements and shifts that occur during the AR in S. ingentis sperm must be 
viewed within the context of this unique system. These cells, after transfer to a female, 
are stored for several weeks or more in exoskeletal seminal receptacles during which 
time they undergo maturational and/or capacitational changes (Clark el ai, 1984). 
During storage they are separated from the seawater (pH ca. 8.0-8.2) by only the 
seminal plasm in which they are embedded. Thus, these cells probably maintain a 
pH; in the same region as that found in their environment (seawater). At least two 
possibilities arise that would functionally explain why these sperm possess such a high 
unactivated pH,: ( 1 ) the energetic costs of maintaining an elevated pH s are less than 
if pHj were depressed below physiological levels (ca. 7.0-8.0); or (2) since sperm un- 
dergo maturational/capacitational changes while in the seminal receptacles of the 
female, the elevated pH ; might be associated with these processes (e.g., in the preven- 
tion of premature filament formation). These, of course, are not all inclusive nor are 
they mutually exclusive; rather, they are questions that await investigation. 

ACKNOWLEDGMENTS 

We thank C. Hand, J. Shenker, and R. Nuccitelli for their critical comments and 
discussion. This work was supported by grants from Sea Grant (NA85AA-D-SG140 
R/A-6 1 ) to WHC, and Sea Grant (NA85 AA-D-SG 1 40 R/A 62) and NSF (DMB85- 
18194)toJHCandLMC. 

LITERATURE CITED 

ANDERSON, S. L., W. H. CLARK JR., AND E. S. CHANG. 1985. Multiple spawning and molt synchrony in 
a free spawning shrimp (Sicyonia ingentis: Penaeoidea). Biol. Bull. 168: 377-394. 

AUSTIN, C. R. 1968. Ultrastructure of Fertilization. Holt, Rinehart and Winston, New York. 145 pp. 

BUSA, W. B., AND R. NUCCITELLI. 1984. Metabolic regulation via intracellular pH. Am. J. Phvsiol. 246: 
R409-R438. 

CAVANAUGH, G. M. ed. 1956. Formulae and Methods of the Marine Biological Laboratory Chemical 
Room. Woods Hole. Pp. 55-56. 

CLARK, W. H., JR., M. G. KLEVE, AND A. I. YUDIN. 1981. An acrosome reaction in natantian sperm. J. 
Exp. Zool. 218:279-291. 

CLARK, W. H., JR., A. I. YUDIN, F. J. GRIFFIN, AND K. SHIGEKAWA. 1984. The control of gamete activa- 
tion and fertilization in the marine Penaeidae, Sicyonia ingentis. Pp. 459-472 in Advances in 
Invertebrate Reproduction 3, W. Engels et a/., eds. 

DAN, J. C. 1952. Studies on the acrosome reaction. I. Reaction to egg-water and other stimuli. Biol. Bull. 
107: 54-66. 

DAN, J. C. 1967. Acrosome reaction and lysins. Pp. 237-293 in Fertilization, Vol. 1, C. B. Metz and A. 
Monroy, eds. Academic Press, New York. 



A pH, DECREASE AT SPERM ACTIVATION 323 

GRINSTEIN, S., S. COHEN, H. M. LEDERMAN, AND E. W. GELFAND. 1984. The intracellular pH of quies- 
cent and proliferating human and rat thymmic lymphocytes. J. Cell. Physiol. 121: 87-95. 
JOHNSON, R. G., AND A. SCARPA. 1976. Internal pH of isolated chromaffin vesicles. J. Biol. Chem. 251: 

2189-2191. 
KLEVE, M. G., A. I. YUDIN, AND W. H. CLARK JR. 1980. Fine structure of the unistellate sperm of the 

shrimp, Sicyonia ingentis (Natantia). Tissue Cell 12: 29-49. 
LOPO, A. C. 1983. Sperm-egg interactions in invertebrates. Pp. 269-324 in Mechanism and Control of 

Fertilization, J. F. Hartmann, ed. Academic Press, New York. 
LOWRY, O. H., N. J. ROSEBROUGH, A. L. FARR, AND R. J. RANDALL. 1951. Protein measurement with 

the Folin-phenol reagent. J. Biol. Chem. 193: 265-275. 
MEIZEL, S., AND D. W. DEAMER. 1978. The pH of the hamster sperm acrosome. J. Histochem. Cvtochem. 

26:98-105. 
MATSUI, T., I. NISHIYAMA, A. HINO, AND M. HOSHI. 1986. Intracellular pH changes of starfish sperm 

upon the acrosome reaction. Dev. Growth Differ. 28: 359-368. 

PRESSMAN, B. C. 1976. Biological applications of ionophores. Ann. Rev. Biochem. 45: 501-530. 
Roos, A., AND W. F. BORON. 198 1 . Intracellular pH. Physiol. Rev. 61: 296-434. 
SHACKMAN, R. W., R. CHRISTEN, AND B. M. SHAPIRO. 1981. Membrane potential depolarization and 

increased intracellular pH accompany the acrosome reaction of sea urchin sperm. Proc. Natl. 

Acad. Sci. 78: 6066-6070. 
SHAPIRO, B. M., AND E. M. EDDY. 1980. When sperm meets egg: biochemical mechanisms of gamete 

interaction. Int. Rev. Cytol. 66: 257-302. 
SHIGEKAWA, K., AND W. H. CLARK JR. 1986. Spermiogenesis in the marine shrimp, Sicvonia ingentis. 

Dev. Growth Differ. 28: 95-1 12. 
TILNEY, L. G., D. P. KIEHART, C. SARDET, AND M. TiLNEY. 1978. Polymerization of actin iv . Role of 

Ca ++ and H + in the assembly of actin and membrane fusion in the acrosome reaction of echino- 

derm sperm. J. Cell Biol. 77: 536-550. 
WADDELL, W. J., AND T. C. BUTLER. 1959. Calculation of intracellular pH from the distribution of 5,5- 

dimethyl-2,4-oxazolidinedione (DMO). J. Clin. Invest. 38: 720-729. 
WORKING, P. K., AND S. MEIZEL. 1983. Correlation of increased intracrosomal pH with the hamster 

sperm acrosome reaction. J. E.\p. Zoo/. 227: 97-107. 



Reference: Biol. Bull. 173: 324-334. (October, 1987) 



A MORPHOLOGICAL EXAMINATION OF GASTRULATION IN A 
MARINE ATHECATE HYDROZOAN 

VICKI J. MARTIN 

Department of Biological Sciences, University of Notre Dame. Notre Dame, Indiana 46556 

ABSTRACT 

The early embryonic development of the marine hydrozoan Halocordyl disticha 
is examined via light histology and transmission electron microscopy. Particular em- 
phasis is devoted to the gastrula and the mode of gastrulation. Cleavage in Halocordyl 
disticha is irregular, total, and asynchronous resulting in the production of stereoblas- 
tulae. Each stereoblastula forms a blastopore at the future posterior end of the larva 
and gastrulates via invagination to produce a lecithotrophic planula larva. During 
gastrulation spherical surface cells radially migrate toward the blastopore, become 
cuboidal-shaped in the region of the pore, and disappear to the interior of the embryo. 
Gastrulation requires 2 h to complete, during which time the ectoderm becomes sepa- 
rated from the endoderm by a mesoglea, interstitial cells arise in the central endo- 
derm, and the embryo elongates to form a planula larva. This study presents the first 
documented example of invagination in the Hydrozoa. 

INTRODUCTION 

Cnidarians represent an early phase of metazoan evolution. Their simple architec- 
ture combined with their exceptional morphogenetic plasticity and adaptability make 
them popular animals for examining developmental processes and principles. The 
phylum is unusual in that its postembryonic development has been more thoroughly 
investigated than its embryogenesis. This is surprising because the cnidarians offer 
excellent material for the study of the evolution of embryogenesis. In the simpler 
cnidarians embryogenesis may appear "anarchic," whereas in the more advanced 
forms one sees complex mosaic patterns of embryogenesis (Metschnikoff, 1886; 
Carre, 1969). 

Since the work of Metschnikoff in the late 1 80(Ts a few papers dealing with embry- 
onic development of the Hydrozoa have been published (Van de Vyver, 1964, 1967, 
1980; Bodo and Bouillon, 1968; Mergner, 1972; Freeman, 1981; Martin and Archer, 
1986). Mergner (1972) attempted to provide a general overview of the processes in- 
volved in cleavage, germ layer formation, and postembryonic development and con- 
cluded that there was great diversity in hydrozoan developmental modes. Van de 
Vyver (1980) analyzed via light histology modes of cleavage, germ layer formation, 
and postembryonic development of several species of hydrozoans and concluded that 
the modes of embryonic development in the Hydrozoa are restricted to a very few 
which are commonly distributed in the animal kingdom. She stated that two types of 
cleavage commonly occur in the Hydrozoa and that cleavage is dependent upon yolk 
quantity of the egg. As a general rule, cleavage for eggs adequately supplied with 
yolk is radial, total, and adequal. Such cleavage is characteristic of eggs of Filifera or 

Received 21 April 1987; accepted 27 July 1987. 



324 



GASTRULATION IN A HYDROZOAN 325 

Capitata corynoidea developing in a gonophore or spawned into the water by free- 
swimming medusae. Large yolk-filled eggs such as those of Capitata tubularoidea 
undergo irregular cleavage. Van de Vyver (1980) further concluded that the most 
important difference between the types of cleavage in the hydrozoans is the presence 
or absence of a blastocoele since its occurrence will determine the mode of germ 
layer formation. MetschnikorT( 1886) proposed that eggs released by free-swimming 
medusae form coeloblastulae while others developing inside gonophores form ste- 
reoblastulae. Van de Vyver (1980) suggested that although Metschnikoffs first point 
might be true, his second is certainly not. Furthermore, Van de Vyver (1980) stated 
that within the Hydrozoa the processes of gastrulation are numerous and may vary 
from species to species. Within the Hydrozoa gastrulation has been shown to occur 
via either ingression, multipolar ingression, delamination, or simple cellular rear- 
rangements (Jagersten, 1972; Tardent, 1978: Van de Vyver, 1980). Invagination has 
not yet been reported in the Hydrozoa although it is common in anthozoans and 
scyphozoans (Tardent, 1978; Van de Vyver, 1980). Van de Vyver (1980) reported 
that polar ingression, multipolar ingression, and delamination are characteristic of 
coeloblastulae, whereas in stereoblastulae the cells which occupy the periphery of the 
embryo simply become progressively different from those situated in the center. She 
claimed that no movements of cells occur in stereoblastulae. 

From the above discussion it is quite clear that our basic knowledge concerning 
embryonic morphogenesis in the cnidarians is sketchy and that additional studies of 
embryogenesis in this phylum are needed. In this study the early development of a 
marine athecate hydrozoan, Halocordyl disticha, is analyzed via light histology and 
transmission electron microscopy. Particular emphasis is devoted to the gastrula. Ha- 
locordyl disticha is a member of the suborder Capitata and forms free-swimming 
medusae which release eggs and sperm into seawater where fertilization is external. 
Cleavage is irregular, asynchronous, and total resulting in the formation of stereoblas- 
tulae which gastrulate via invagination to produce lecithotrophic planula larvae. This 
study presents the first documented example of invagination in the Hydrozoa. 

MATERIALS AND METHODS 

Mature colonies of the marine hydrozoan Halocordyl disticha were collected from 
pier pilings at the University of North Carolina Institute of Marine Sciences in More- 
head City, North Carolina. Fronds from mature male and female colonies were 
placed together in large finger bowls of filtered seawater. Subsequently, the bowls 
were placed in the dark at 6:00 pm and returned to the light at 9:00 pm. Within 1 
hour after exposure to light early cleavage stages were found in the bottoms of the 
dishes. These embryos were transferred to small finger bowls of filtered seawater and 
reared at 23C to the planula stage. 

Early cleavage embryos, late cleavage embryos, blastulae, gastrulae, and young 
planulae were prepared for either light histology or transmission electron microscopy. 
Animals for light microscopy were fixed for 1 hour in 10% formalin in seawater, 
dehydrated in an ethanol series, and embedded in Paraplast Plus paraffin. Serial sec- 
tions, 10 nm thick, were mounted on glass slides and stained with either Azure B or 
the SchifFs reagent (nucleal feulgen reaction). Live embryos and prepared histological 
sections were photographed with a Zeiss standard research microscope. Embryos un- 
dergoing cleavage and gastrulation were also continuously examined under the mi- 
croscope until young planulae were formed. 

Samples for electron microscopy were fixed for 1 h in 2.5% glutaraldehyde, pH 



326 V. J. MARTIN 

7.4, in 0.2 M phosphate buffer. They were postfixed for 1 h in 2% osmium tetroxide, 
pH 7.2, in 1.25% sodium bicarbonate. Specimens were dehydrated in an ethanol 
series, infiltrated, and embedded in Spurr's embedding media. Blocks were serially 
secti. ..',' on a Porter-Blum MT-2B ultramicrotome, placed on 150-mesh copper 
grid , and stained with 3.5% uranyl acetate in ethanol followed by lead hydroxide. 

ids were examined and photographed with a Hitachi H-600 transmission electron 
microscope. 

Surface cells of live embryos at various stages of gastrulation were marked with 
Nile Blue and subsequently monitored for their movement. The marking technique 
involved using a 0.01% solution of Nile Blue in seawater. The dye was drawn into 
microcapillary pipettes with varying bore diameters. Embryos were immobilized for 
marking by placing them in a tiny groove in the bottom of a Falcon small plastic petri 
dish. The dye-containing micropipettes were gently touched to the surfaces of the 
embryos for 30-60 seconds producing small blue patches of marked cells of varying 
diameter (depending upon pipette bore size) along the animal surface. Previous 
marking studies using planulae indicate that Nile Blue is nontoxic at the 0.01% con- 
centration and embryos stained with Nile Blue retain the dye for several days. The 
dye will not diffuse into unstained tissue (Martin, unpub.). After marking the gastrula 
cells, half of the animals were removed from the grooves and returned to small dishes 
containing filtered seawater. The other half were left immobilized in the grooves and 
their dishes placed in a moist chamber to prevent samples from drying. The marked 
cells of the immobilized and free-moving animals were continuously examined 
throughout gastrulation for change in axial position. 

RESULTS 

Cleavage in embryos ofHalocordyldisticha is holoblastic, unequal, and asynchro- 
nous (Figs. 1-4). Cleavage begins 1 h after fertilization and results in the formation 
of blastomeres of unequal size. A period of early cleavage extends to the beginning 
of 6 h postfertilization during which time no one embryo cleaves in exactly the same 
fashion. Such embryos assume numerous bizarre shapes and sizes and reach the 128- 
256 cell stage (Martin and Archer, 1986). Early cleavage is rapid and by 6 to 8 h 
postfertilization a stereoblastula (late cleavage) is formed (Figs. 3, 4). The stereoblas- 
tula assumes the shape of a sphere and the blastomeres are more uniform in size than 
during early cleavage. The stereoblastula is ca. 230 nm in diameter and consists of an 
outer layer of small spherical blastomeres surrounding an inner layer of larger spheri- 
cal blastomeres (Fig. 4). 

By 8 h postfertilization the surface of the embryo is smooth and a single indenta- 
tion appears at one pole of the embryo (Figs. 5, 6, 10-12). This indentation corre- 
sponds to a blastopore, and the pole at which it forms marks the future posterior pole 
of the planula (Figs. 7, 8, 13, 14). This stage represents gastrulation and the young 
gastrula is ca. 250 p.m long and 190 ^m wide (Fig. 1 1). Gastrulation requires 2 hours 
to complete and during this time a number of events occur (Figs. 5-20). The initial 
blastopore indentation will deepen to form a groove (Figs. 5, 6, 10-12). Some of 
the cells on the surface migrate in a radial fashion toward the deepening blastopore, 
invaginate over the lips of the pore, and disappear to the inside. Such movement of 
cells is easily visualized using the Nile Blue marking procedure. Marked patches of 
blue cells move toward the blastopore, briefly inhabit the lips of the pore, and eventu- 
ally disappear from the surface of the animal. Hence on a marked animal a blue patch 
of cells can be traced from the surface to the blastopore region, and ultimately to the 



GASTRULATION IN A HYDROZOAN 



327 







B-J 



EN 




B- ? 
{ 



4 



m 



B- 





6 




FIGURES 1 -9. Histological sections of developing embryos of Halocordyl disticha. 
FIGURE 1 . Early cleavage embryo (3 h postfertilization) X200. 

Early cleavage embryo (3 h postfertilization) X200. 

Late cleavage 7-h embryo (stereoblastula). Note the absence of a mesoglea and interstitial 



FIGURE 2. 
FIGURE 3. 
cells. X200. 
FIGURE 4. 
FIGURE 5. 



Stereoblastula (7 h postfertilization) X200. 

Early 8-h gastrula. An early blastopore (B) is visible. Separation of the two germ layers is 
apparent and interstitial cells (arrow) appear in the central endoderm. E, ectoderm; EN, endoderm. X200. 

FIGURE 6. Mid-gastrula stage. The blastopore (B) has deepened to form a groove. A mesoglea (black 
arrow) is seen as are numerous interstitial cells (white arrow). X200. 

FIGURE 7. Nine-hour gastrula which has begun to elongate. The blastopore (B) is located at the 
posterior pole of the embryo and a mesoglea is present (black arrow). Cells migrating over the lips of the 
blastopore to the inside are visible (white arrows). X200. 

FIGURE 8. Elongating 9-h gastrula. Central endodermal interstitial cells (white arrow) are distin- 
guishable from the outer endodermal gastrodermal cells (black arrow). B, blastopore. X200. 

FIGURE 9. Ten-hour planula. The ectoderm is separated from the endoderm by an acellular meso- 
glea (M). White arrow, interstitial cells; Black arrow, gastrodermal cells. X200. 



328 



V. J. MARTIN 




10 



11 





12 



13 





15 



FIGURES 10-15. Gastrulation in Halocordyl disticha. 

FIGURE 10. Early 8-h gastrula with a slight indentation (blastopore) at the future posterior pole. X75. 

FIGURE 1 1 . Mid-8-h gastrula with a deepening blastopore. x75. 

FIGURE 1 2. Late 8-h gastrula with a prominent blastopore. Lips of the blastopore are visible and the 
embryo has begun to elongate. X75. 

FIGURE 1 3. Elongating 9-h gastrula. The embryo has a distinct anterior end and a posterior end. The 
blastopore is visible at the posterior end. X75. 

FIGURE 14. Late 9-10-h gastrula which has elongated. The blastopore is still visible at the posterior 
pole. X75. 

FIGURE 15. Ten-hour planula. The blastopore has completely closed producing a 2 germ layered 
planula larva. X75. 



GASTRULATION IN A HYDROZOAN 



329 



B 




FIGURE 1 6. Longitudinal section through the blastopore region of an 8-h embryo. As cells move into 
the region of the pore they change from a spherical shape to a cuboidal shape (arrows). Microvilli and cilia 
from cells forming the lips of the blastopore project into the space of the pore (B). X4900. 



animal interior. Time required for such patch movement (i.e., from the initial 
marked surface position to the disappearance at the blastopore) varies anywhere from 
1 5-30 minutes. The shape of the migrating cells changes as they move inward. 

Examination of the blastopore region via transmission electron microscopy illus- 



330 



V. J. MARTIN 




FIGURE 17. Enlargement of a portion of the blastopore region of an 8-h embryo. Cuboidal-shaped 
cells form the lips of the pore. B, groove of the blastopore; N, nuclei of cells in the blastopore area. X5000. 



trates the true nature of the indentation (Figs. 16-18). In the region of the blastopore 
groove, spherical surface cells become cuboidal (Figs. 16, 17). Hence the cells forming 
the lips of the blastopore are cuboidal. Such cuboidal-shaped cells possess cilia and 
microvilli that project into the groove of the pore (Figs. 16-18). The cuboidal cells of 
the blastopore eventually disappear to the interior of the embryo. As cells invaginate 
a clear separation of the ectoderm and endoderm becomes visible with the formation 
of an acellular mesoglea (Figs. 5-8). 

During gastrulation there is localization of embryonic tissue types within the en- 
doderm. The presumptive gastrodermal cells become distinguishable from the mes- 
enchymal-like interstitial cells (Figs. 5-8, 19, 20). The interstitial cells appear as an 
aggregate of cells in the central endodermal core of the embryo during invagination. 
At this time some cytodifferentiation has begun since interstitial cells stain more 
darkly with azure B than do the more peripheral gastrodermal cells (Fig. 8). In the 
early gastrula (just prior to blastopore formation) the central blastomeres consist of 
large yolk-filled masses (Fig. 19). Such blastomeres appear to be loosely packed in 
the center of the embryo as indicated by the large intercellular spaces between the 
blastomeres (Fig. 19). At this stage interstitial cells are not yet present. Once invagina- 
tion begins the loose arrangement of the central blastomeres is lost (intercellular 
spaces disappear) and clusters of small round interstitial cells appear in the center of 
the embryo (Fig. 20). Such interstitial cells become clearly segregated from the outer 
forming columnar gastrodermal cells. 

Between 8 and 10 h postfertilization the gastrula elongates in an anterior-posterior 



GASTRULATION IN A HYDROZOAN 



33 







FIGURE 18. Cross-section through the blastopore region of an 8-h embryo. Microvilli and cilia of 
migrating cells extend into the space of the blastopore (B). N, nucleus of cell in region of the blastopore. 
X6700. 



direction to form a young planula (Figs. 7-9, 12-15). The 10-h planula is ca. 350 
long, 180 /urn wide in the anterior region, 170 ^m wide in the mid region, and 120 
^um wide in the tail (Fig. 15). By 10 h the planula has a distinct anterior end and 
posterior end. The blastopore is located at the posterior pole of the planula and will 
soon close (Figs. 14, 15). No gastrovascular cavity or mouth is found in the planula 
at any stage of its development. The 10-h planula will elongate to form a mature 
planula (24-96 h old depending on temperature) which will attach via its anterior 
end to a substrate and undergo metamorphosis. 

. 

DISCUSSION 

Within the Cnidaria the processes of gastrulation are numerous and vary widely 
from species to species (Tardent, 1 978). Among the anthozoans gastrulation has been 
reported to occur via either invagination, delamination, multipolar ingression, or a 
combination of invagination and polar ingression (Tardent, 1978). In scyphozoans 
gastrulation may occur via invagination, polar ingression, multipolar ingression, or 
invagination plus polar immigration (Tardent, 1978). In hydrozoans examples of 
gastrulation by polar ingression, multipolar ingression, and delamination have been 
reported. However, until now no examples of invagination have been documented 
(Jagersten, 1972; Tardent, 1978). 

Jagersten (1972) provided a brief overview of gastrulation in the cnidarians and 
stated that within the phylum a connection existed between the mode of gastrulation 
and whether the formed larva was lecithotrophic or planktotrophic. In species which 
gastrulate via either delamination, multipolar ingression, or unipolar ingression, the 



332 



V. J. MARTIN 




FIGURE 1 9. Central inner blastomeres of an early 8-h embryo. These central endoblast cells are filled 
with yolk and are separated from each other by large intercellular spaces. No distinguishable interstitial 
cells are yet present. N, nucleus of central endoblast cell. X4000. 

FIGURE 20. Central endoblast region of a 9-h embryo. Clusters of young interstitial cells are visible. 
As the interstitial cells increase in number the intercellular space decreases and the central region of the 
embryo assumes a more compact appearance. The interstitial cells are completely set apart from the outer 
gastrodermal cells during gastrulation. X4000. 



GASTRULATION IN A HYDROZOAN 333 

derived larvae exhibit lecithotrophy and never planktotrophy. In species which pro- 
duce actively feeding larvae (planktotrophic) the mode of gastrulation is via invagina- 
tion. Jagersten (1955, 1959) presented arguments supporting the ideas that the origi- 
nal method of gastrulation in the cnidarians was via invagination, that the plankto- 
trophic larval life was the primitive condition, and that lecithotrophy was a secondary 
trait which arose independently on different occasions within the phylum. Further- 
more he stated that lecithotrophy is dominant among the Cnidaria. Jagersten (1972) 
and Widersten (1968) proposed that the most primitive features of the phylum are 
found within anthozoans and the most altered within the hydrozoans. Jagersten 
(1972) further said that lecithotrophy may occur in larvae which exhibit invagination 
(e.g., PachycerinatKus). Despite the moderate quantity of yolk in the eggs of these 
animals, invagination persists. 

This study documents the occurrence of invagination in the Hydrozoa. Embryos 
ofHalocordyl disticha form stereoblastulae which gastrulate via invagination to pro- 
duce lecithotrophic planula larvae. Marking studies clearly indicate that surface cells 
migrate to the blastopore, occupy the lips of the pore, and eventually disappear to the 
interior of the gastrula. Neither a mouth nor a gastrovascular cavity form in these 
planulae. The absence of a mouth in cnidarian embryos which gastrulate via invagi- 
nation is not uncommon, as examples also exist among the scyphozoans (Amelia, 
Cyanea) (Jagersten, 1972). 

Jagersten (1972) claimed that the common ancestor of the Metazoa included a 
Gastrea form, a creature with both an alimentary cavity and a mouth. He proposed 
that the almost universal distribution of the invagination gastrula was conclusive evi- 
dence for the Gastrea theory. The hydrozoans can now be added to this universal list 
as invagination gastrulae are found within this class. Furthermore, if Widersten 
(1968) and Jagersten (1972) are correct in their assumptions that invagination is the 
primitive condition within the Cnidaria and that the anthozoans are the more primi- 
tive class, then the invagination process as described in this paper for a marine hydro- 
zoan may illustrate a stubborn retention of this original primitive condition. Clearly, 
further investigations of early embryogenesis in the Hydrozoa concentrating on 
modes of gastrulation are needed to complement the work presented for Halocordyl 
disticha. 

ACKNOWLEDGMENTS 

I thank Margaret Martin for her help in collecting animals and William Archer 
for his technical assistance. This research was supported in part by a grant from the 
National Science Foundation, DCB-8702212. 

LITERATURE CITED 

BODO. F., AND J. BOUILLON. 1968. Etude histologique du development de quelques Hydromeduses de 

Roscoff: Phialidium hemisphaencum (L.). Obelia sp. Peron et Lesieur, Sarsia eximia (Sars), Gon- 

ionemus vertens Aqassiz. Cah. Biol. Afar. 9: 69-104. 
CARRE, D. 1969. Etude du developpement larvaire de Sphaeronectes gracilis (Claus. 1873) et de Sphaero- 

nectes irregularis (Claus, 1873), Siphonophores Calycophores. Cah. Biol. Mar. 10: 3 1-34. 
FREEMAN, G. 1981. The role of polarity in the development of the hydrozoan planula larva. Roux'sArch. 

Dev.Biol. 190: 168-184. 
JAGERSTEN, G. 1955. On the early phylogeny of the Metazoa. The Bilaterogastrea-theory. Zool. Bidr. 

Upps. 30:321-354. 
JAGERSTEN, G. 1959. Further remarks on the early phylogeny of the Metazoa. Zool. Bidr. Upps. 33: 79- 

108. 



334 V. J. MARTIN 

JAGERSTEN, G. 1972. Cnidaria. Pp. 13-22 in Evolution of the Metazoan Life Cycle A Comprehensive 

Theorv. Academic Press, New York. 
MARTIN, V., AND W. ARCHER. 1986. A scanning electron microscopic study of embryonic development 

of a marine hydrozoan. Biol. Bull. 171: 1 16-125. 
MERGNER, HL 1972. Pp. 1-84 in Experimental Embryology of Marine and Freshwater Invertebrates. 

North-Holland, Amsterdam. 
ME is i i.NiKOFF, E. 1 886. Embryologische studien an Medusen. Ein Beit rag zur GenealogiederPrimitivor- 

gane. Alfred Holder, Vienna. 
TARDENT, P. 1978. Coelenterata, Cnidaria. Pp. 199-302 in Morphogenese der Tiere. Gustav Fischer Ver- 

lag, Stuttgart. 
VAN DE VYVER, G. 1964. Etude histologique du developpement embryonnaire d'Hydractinia echinata 

(Flem.). Cah. Biol. Mar. 5: 295-310. 
VAN DE VYVER, G. 1967. Etude du developpement embryonnaire des hydraires athecates (gymnoblas- 

tiques) a gonophores. I. Formes a planula. Arch. Biol. 78:451-518. 
VAN DE VYVER, G. 1980. A comparative study of the embryonic development of Hydrozoa athecata. Pp. 

109-120 in Developmental and Cellular Biology of Coelenterates, P. Tardent and R. Tardent, 

eds., Elsevier/North-Holland, New York. 
WIDERSTEN, B. 1968. On the morphology and development in some cnidarian larvae. Zoo/. Bidr. Upps. 

37: 139-182. 



Reference: Biol. Bull. 173: 335-344. (October, 1987) 



VARIABILITY IN THE PATTERN OF SEXUAL REPRODUCTION OF 
THE CORAL STYLOPHOR.4 PISTILLATA AT EILAT, RED SEA: 

A LONG-TERM STUDY 

B. RINKEVICH* AND Y. LOYA 

Department of Zoology. The George S. Wise Faculty of Life Sciences, Tel Aviv University, 

Ramat Aviv 69978, Israel 

ABSTRACT 

Sexual reproduction of the Red Sea coral Stylophora pistillata was followed at 
Eilat in a long-term study (1974-1984). Field examination of over 9000 colonies 
through 1 19 months indicated that S. pistillata had a reproductive season of approxi- 
mately 8 months (varying from 6 to 9 months). Premature planulae and eggs were 
aborted following winter storms, resulting in a lowering of the planular index and the 
number of female gonads per polyp. Histological examinations of tissue from 20 large 
colonies which were studied for several years, until they were found dead in situ, 
indicated that either sexuality (reproductive states) and/or fecundity could be com- 
pletely altered from one reproductive season to the next: i.e., hermaphroditic colonies 
exhibiting high fecundity in one season became male or even sterile thereafter, and 
vice versa. In addition, great variability in reproduction between successive years was 
recorded in sexuality and in the fecundity of shallow water populations. Shallow wa- 
ter colonies (5 m) possessed up to 5 times more female gonads per polyp and shed 5 
to 20 times more planulae than deep water colonies (25 to 45 m) in which the repro- 
ductive season is 2 to 3 months shorter. 

We suggest that the changes in the hermaphroditic, male, or sterile modes of re- 
production in S. pistillata are from energy limitations and stress conditions. Since 
reproductive activity probably involves significant energetic expenditures, any stress 
or diminution in energy resources affects sexuality or fecundity. This should be con- 
sidered before formulating any general hypothesis on coral reproduction. 

INTRODUCTION 

Much information concerning reproductive biology of scleractinian corals has 
recently become available. Fadlallah (1983) reviewed past studies and provided a list 
of almost 90 species in which several known reproductive characteristics are pre- 
sented. More recent studies (Harriott, 1983; Szmant-Froelich et ai. 1984; Shlesinger 
and Loya, 1985; Wallace, 1985; Willis et ai., 1985; Babcock et al, 1986; Szmant, 
1986) provide information on reproductive patterns of more than 100 additional 
species of corals. 

Although this list of studied species is impressive, data on scleractinian reproduc- 
tion is still scanty, especially that dealing with their reproductive ecology. These stud- 
ies evaluated sizes, shapes, and numbers of gonads, and attempted to establish repro- 
ductive seasonality, lunar periodicity, mode of reproduction, planula characteriza- 
tions, and behavior. However, most studies were based on observations and 

Received 30 January 1987; accepted 31 July 1987. 

* Present Address: Hopkins Marine Station of Stanford University, Pacific Grove, California 93950. 

335 



336 B. RINKEVICH AND Y. LOYA 

experiments carried out within a period of a year or less. Only a few studies dealt 
with longer periods ranging from two (Atoda, 1947a, b; Harriott, 1983; Jokiel, 1985; 
Wallace. ' 985 ) to three years (Kojis and Quinn, 198 la; van Moorsel, 1983; Stoddard 
and Black, 1985). Consequently, studies on sexual reproduction often fall short in 
documenting many aspects of coral reproduction (Fadlallah, 1983). Detailed infor- 
mation on coral reproduction could clarify many aspects of their life history patterns 
and provide a better understanding of the coral reef as a whole. 

Stylophora pistillata (Esper) is one of the most abundant coral species in the Gulf 
of Eilat, Red Sea. Some aspects of the reproduction of this species have already been 
studied in the field and the laboratory. Descriptions of planulae and gonads have 
been made (Rinkevich and Loya, 1979a). In addition, synchronization in breeding, 
colony size in relation to fecundity, onset of reproduction, reproduction within a 
single colony, and seasonality of planulation were also reported (Rinkevich and Loya, 
1979b). This paper summarizes results of a ten-year study on the reproduction of S. 
pistillata which elucidate some general conclusions characterizing coral reproductive 
activities. 

MATERIALS AND METHODS 

Reproductive activity of S. pistillata was studied from March 1974 to January 
1984 (most intensively from 1976 to 1980). The study area was located in front of 
the H. Steinitz, Marine Biological Laboratory at Eilat, Gulf of Eilat, Red Sea, and 
was visited regularly once a month during the ten-year study period. Large colonies 
(mean geometric radius, 7 > 20 cm) were sampled from both shallow (3-5 m) and 
deep water (25-60 m) populations. 

Reproduction was studied by two techniques: collections of shed planulae (see 
below) and examinations of gonads in histological sections (Rinkevich and Loya, 
1979a, b). A single branch was sampled from each colony. This branch represents the 
reproductive state of the entire colony (Rinkevich and Loya, 1979b). The number of 
female gonads was counted within serial sections for each tested polyp and quantita- 
tive data were obtained on the average number of eggs per polyp in a given specimen 
(6-18 polyps per sample). Male gonads were not counted because of the difficulty of 
following them in serial histological sections as a result of the irregular shape of a 
typical male gonad (Rinkevich and Loya, 1979a). Tissue samples were always taken 
near the bases of branches since few polyps from the tips contain genital cells (Rinkev- 
ich and Loya, 1979b; Kojis and Quinn, 198 la). 

Early in the study, planulae were collected in situ by covering large colonies with 
plankton nets in the late afternoon and removing the nets at midnight (Rinkevich 
and Loya, 1979a). However, due to the difficulties with this technique during night 
diving (especially with the deep-water colonies), planulae were collected from coral 
branches that were brought into the laboratory. The branches were carefully removed 
underwater using wire cutters, and transported to the laboratory within 30 min after 
sampling in closed, separate plastic bags. The water in each bag was checked for the 
appearance of planulae. Each sample was put separately in a 5 1 glass aquarium, con- 
taining filtered seawater, and left overnight. Planulae were shed during the night (Rin- 
kevich and Loya, 1 979a). Although handling stimulated planula release, it is assumed 
that these planulae were prepared for shedding. This assumption was supported by 
the finding that the released planulae were fully developed. Since conditions in all 
treated samples were similar, we concluded that collection procedures did not affect 
the results. Planulae were counted by sight and removed by pipettes. The seawater 
was then filtered through a plankton net (100 ^m) and all remaining planulae were 



VARIABILITY IN SEXUAL REPRODUCTION OF A CORAL 



337 



TABLE I 

Some characterizations of recorded southern storms in Eilat 



Waves 



Winds 



Date 


No. of 
storms 


Max. height 
(m) 


Max. length 
(m) 


General 
direction 


Max. speed 
(km/h) 


Feb. 1979 


3 


2 


12 


SE 


nd 


Apr. 1979 


3 


1 + 


nd 


S 


nd 


Nov. 1979 


1 


1 


6 


SE 


20 


Dec. 1979 


2 


2 


nd 


SE 


nd 


Feb. 1980 


2 


2.5 


10 


SSE 


35 


Mar. 1980 


1 


nd 


nd 


nd 


nd 


Apr. 1980 


1 


nd 


nd 


nd 


nd 


Jan. 1981 


2 


1.5 


20 


S, SE, SW 


25 


Feb. 1981 


3 


1.2 


27 


SSE 


18 


Mar. 1981 


1 


0.5 


nd 


S.SE 


nd 



nd = no available data. (Personal communication, C. Porter, Israel Oceanographic and Limnological 
Research Ltd., Eilat). 



collected and counted. Sampled branches were placed on a filter paper for 15 min to 
remove excess water and weighed (accuracy to the nearest 1 g). In most cases branch 
weights ranged between 100-200 g. Results are presented as number of released plan- 
ulae per 100 g of coral skeleton, during one night. 

In addition, the release of planulae from mature colonies ( r > 20 cm) was checked 
each month in situ where several branches were carefully broken from many colonies. 
This procedures stimulated the release of planulae in colonies which were in a repro- 
ductive state. The shed planulae were easily seen and traced by sight underwater. A 
planular index was then formulated (see below), which took into account the relative 
number of planulae shed and the percentage of reproducing colonies. Since variabil- 
ity in the fecundity between different colonies within the population was high (Rin- 
kevich and Loya, 1979b), up to 90 large colonies were sampled each month (in 2-3 
replicates, at least 30 colonies in each) to assess the validity of the planular index. The 
index sign (-) was given when none of the sampled colonies released any planulae; 
(H ) when very few planulae were released (total number of 1-5 planulae from the 
30 tested colonies in each replicate); (+) when about one third of the colonies released 
few planulae; (++) when up to two thirds of the colonies shed planulae (many or few; 
many = any small fragment broken from the colony released about two planulae); 
(+++) when most or all of the colonies shed planulae. 

The most severe storms in the sea at Eilat are known as southern storms, which 
occur during the winter and spring. Some physical parameters of these storms are 
partly documented from February 1979 (Table I). 

RESULTS 
Long-term study on seasonally of planulae shedding 

Plantation in S. pistillata was continuously studied between March 1974 to Jan- 
uary 1 984 by sampling more than 9000 colonies (Table II). The two questions investi- 
gated were whether plantation occurs in the same months from one year to the next 
and how the planular index in the shallow water population fluctuated during the 



338 B. RINKEVICH AND Y. LOYA 



TABLE II 

Monthly planular index in shallow water populations of Stylophora pistillata 
during 119 months of observations 



Planular index* in 



Year Jan. Feb. Mar. Apr. May Jun. Jul. Aug. Sep. Oct. Nov. Dec. 

1974 
1975 
1976 
1977 
1978 
1979 
1980 
1981 
1982 
1983 
1984 

* (-), No planulae; (H ), very few; ( + ), few; ( + + ), intermediate; (+++), large numbers. 



119 months of observations. S. pistillata has a long reproductive season (planulae 
release) lasting approximately 8 months, from December to July (Table II). However, 
the reproductive season ranged 6 months (in 1976) to 9 months (in 1975). In the 
three-month period from August to October, no planulation was ever recorded. Only 
once during the ten-year investigation were very few planulae observed in November 
(in 1975). A marked variation was noted in the December-January-February index 
between different years. Although these months represent the beginning of the repro- 
ductive season (Rinkevich and Loya, 1979b), this variability might also be the result 
of the southern storms which are most severe during the winter (Table I). This phe- 
nomenon is also demonstrated in another part of the present study: in April 1980 a 
southern storm interrupted our field sampling. Branch samples were collected before 
the storm from 10 mature colonies. Nine of them released high numbers of planulae 
(average of 30 28 planulae, per 100 g skeleton, per colony). One day after the storm 
samples were collected for histological study from 1 3 other mature colonies inhabit- 
ing the same area and depth. Only eight colonies contained low numbers of female 
gonads while the others were either sterile or contained only male gonads. The aver- 
age number of female gonads per polyp, per colony was very low (0.4 0.6), much 
lower than other April months (for more detail, see Tables VI and IV, respectively). 

Long-term study on reproductive states 

Two separate sets of experiments followed the long-term state of reproduction in 
shallow water populations. In the first experiment 20 large colonies (7 > 20 cm) were 
chosen (December 1 976) and sampled for histological study two to three times a year 
e.g., in the beginning, the peak and the end of the reproductive season over four 
successive reproductive periods, until the deaths of all of them were recorded (Febru- 
ary 1980). Since synchronization in the reproduction activity exists between branches 
(Rinkevich and Loya, 1979b), only one branch was sampled each time from each 
colony. This sampling procedure did not affect survivorship or reproduction (un- 
pub.). Colony mortality was high (Table III), although colonies were carefully chosen 
on the basis of their healthy state (without dead branches or tissue damage). One and 



VARIABILITY IN SEXUAL REPRODUCTION OF A CORAL 339 

TABLE III 

Reproductive state and average number of female gonads per polyp in Stylophora pistillata 
colonies sampled during Dec. 1976 to Feb. 1980 

Average numbers of female gonads per polyp in 
Coral 
no. Dec. 76 Apr. 77** Dec. 77 Apr. 78 Jun. 78 Dec. 78 Apr. 79 Dec. 79 Feb. 80 



1 


-(10) 


0.7(10)* 


D 








2 


0.2(10) 


0.5(10)* 


D 








3 


0.6(10) 


2.9(10)* 


1.2(6) 


2.6(7)* 


0.7(6)* 


-(10) D 


4 


2.4(10) 


1.4(10)* 


0.2(6) 


+(10) 


D 




5 


0.4(10) 


0.4(10)* 


D 








6 


0.1 (10) 


1.9(10)* 


+(6) 


-(10) 


-(10) 


-(8) D 


7 


1.9(10) 


1.1 (10)* 


0.5(6) 


D 






8 


-(10) 


D 










9 


0.6(10) 


1.1 (10)* 


0.7(7) 


2.2(6)* 


S 




10 


-(10) 


0.4(10)* 


+(6) 


+(6) 


-(8) 


1.4(7) 0.4(7) -(9) D 


11 


-(10) 


D 










12 


-(10) 


D 










13 


-(10) 


2.5(10)* 


2.1(10) 


2.5(10)* 


-(6) 


1.8(6) 1.0(8) D 


14 


-(10) 


0.5(10)* 


-(5) 


1-9(11) 


-(10) 


-(10) -(10) D 


15 


-(10) 


1.6(10)* 


1.0(5) 


S 






16 


1.1(10) 


1.3(10)* 


2.3(10) 


S 






17 


-(10) 


D 










18 


-(10) 


0.4(10)* 


D 








19 


-(10) 


0.9(10)* 


-(11) 


3.3(7) 


S 




20 


0.2(10) 


0.9(10)* 


+(7) 


2.1(8) 


-(10) 


1.2(6) D 



December and February months refer to the beginning of the reproductive season, April months to 
the peak of reproduction, and June to the decline phase of the reproductive season (numbers in parentheses 
refer to the number of polyps examined). 

-, Sterile colony; +, Only male gonads present; D, The colony was found dead; *, Planulae detected 
in histological sections; **, Planulae were found in all plankton nets put on marked colonies; S, Destroyed 
by storm. 



two years after the beginning of the study, 60% and 30%, respectively, of the colonies 
were alive. Only one colony of the 20 samples (5%) remained alive after 3 years (Table 
III). A decrease in fecundity was repeatedly observed several months prior to the 
natural death of many of the colonies. In four out of the six dead colonies following 
a period of high fecundity (colonies 9, 15, 16, 19; Table III), the death was attributed 
to southern storms. None of the dead colonies during the first 4 months of the study 
(colonies 8, 11, 12, 17; Table III) contained any genital cells when first sampled. 

The results (Table III) also indicate variability in sexuality (reproductive states: 
male, hermaphrodite, or sterile modes of reproduction) and fecundity of a specific 
colony in different years. Hermaphroditic colonies which exhibit high fecundity in 
one reproductive season may differ in the following reproductive season in which 
they become sterile (colonies 6, 14; Table III), or male (colonies 4, 10; Table III) and 
vice versa. The changes in colony reproductive patterns are further demonstrated in 
colonies sampled in three to four consecutive December months (colonies 3, 6, 10, 
13, 14, 20; Table III). Sexuality or fecundity of five out of these six colonies were 
altered in each December month. 

In the second set of experiments (Table IV), 1 55 large shallow water colonies were 
sampled over 10 successive sampling periods at the beginning and during the peak of 
5 reproductive seasons (April 1 976-April 1 980). Changes among the different seasons 



340 



B. RINKEVICH AND Y. LOYA 



Stylophora 


TABLE IV 
pistillata: reproductive states of shallow water populations 






Colony reproductive state (%) 


A f 1 J 




No. of 








Average temale gonads 


Date 


colonies 


Hermaphrodites 


Males only 


Sterile 


(polyp" 1 colony"') 


Apr. 76 


17 


94 


6 





1.5 0.9 


Dec. 76 


20 


45 





55 


0.4 0.7 


Apr. 77 


16 


100 








1.2 0.8 


Dec. 77 


12 


67 


25 


8 


0.7 0.8 


Apr. 78 


9 


67 


22 


11 


1.6 1.3 


Jun. 78 


16 


31 





69 


0.2 0.4 


Dec. 78 


14 


50 





50 


0.5 + 0.6 


Mar. 79 


26 


88 





12 


1.5 1.1 


Dec. 79 


12 


50 


25 


25 


0.5 + 0.7 


Apr. 80 


13 


62 





38 


0.4 0.6 



either in sexuality or fecundity were observed. For example, each one of the 4 differ- 
ent December months (years 1976, 1977, 1978, 1979) represented different patterns 
of reproductive states: 8-56% of the colonies were sterile, 0-25% males and 45-67% 
were hermaphrodites among the different December months. The same pattern was 
recorded for sexuality of March to April months: 0-38%, 0-22% and 62-100%, re- 
spectively (Table IV). It is concluded that "one year of sampling" is not enough for 
the characterization of reproductive states in this species. 

Reproduction in .shallow versus deep water populations 

Possible differences in reproduction between shallow and deep water populations 
were tested in two sets of experiments. In the first, we analyzed serial histological 
sections of 90 shallow water colonies (from Table IV). The results were compared to 
those of 77 deep water colonies (25-45 m, Table V) sampled on the same days 
during three successive reproductive seasons (April 1978-April 1980). Shallow water 
colonies possessed up to 5 times more female gonads per polyp per colony than deep 
water colonies (P < 0.01, Wilcoxon's signed rank test; Sokal and Rohlf, 1981). This 
phenomenon was most clear during the peak of the reproductive season, March to 



Stylophora pistillata: 


TABLE V 

reproductive states of deep water colonies 








Colony reproductive state (%) 






Depth 


No. of 








Average temale gonads 


Date 


(m) 


colonies 


Hermaphrodites 


Males only 


Sterile 


(polyp" 1 colony" 1 ) 


Mar. 78 


60 


1 


100 








0.7 


Apr. 78 


25-30 


15 


40 


40 


20 


0.3 0.4 


Jun. 78 


25 


15 


7 





93 


0.0 


Dec. 78 


27-30 


15 


86 


7 


7 


0.7 0.6 


Mar. 79 


40-45 


11 


36 


36 


28 


0.4 0.8 


Dec. 79 


27-30 


9 








100 





Apr. 80 


25-30 


12 


8 


50 


42 


0.0 0.1 



VARIABILITY IN SEXUAL REPRODUCTION OF A CORAL 341 

TABLE VI 
Shedplanulae in sample branches of shallow and deep water Stylophora pistillata colonies 



Shallow 



Deep 









Colonies 
which 


Average 
no. of 






Colonies 
which 


Average 
no. of 








shed 


planulae 






shed 


planulae 




Depth 


No. of 


planulae 


(per 100 g 


Depth 


No. of 


planulae 


(per 100 g 


Date 


(m) 


colonies 


(%) 


skeleton) 


(m) 


colonies 


(%) 


skeleton) 


Jan. 79 


3-6 


5 


80 


32 49 


27 


6 








Feb. 79 


3-6 


6 


100 


85 95 


27-30 


4 


25 


1 1 


Mar. 79 


5 


6 


100 


22 20 


40-42 


5 


20 


0.4 1 


Jun. 79 


5 


4 


100 


31 22 


34 


5 


20 


2 5 


Dec. 79 


5 


4 








30 


3 








Jan. 80 


3-5 


9 


89 


1431 


39-42 


9 


33 


4 10 


Feb. 80 


5 


6 


83 


4 2 


30 


6 








Apr. 80 


3-6 


10 


90 


30 28 


25-27 


9 


44 


3 6 


May 80 


3-8 


10 


90 


44 47 


27 


10 


70 


4 7 


Jun. 80 


6-9 


10 


100 


14 10 


27-30 


10 


30 


3 7 


Jul. 80 


4-6 


10 


70 


3 6 


30 


10 


20 


0.2+ 0.4 


Jan. 81 


3-5 


5 


60 


4 8 


30-35 


5 








Feb. 81 


3-5 


6 


67 


5 3 


27-30 


6 


33 


1 1 



April (the average female gonad per polyp in April 1980 is lower than other April 
months because samples were taken immediately after a storm). 

In the second set of experiments, planulae were collected in the laboratory from 
branch samples of 91 shallow water and 88 deep water colonies, on 13 collecting 
dates (Table VI). Significantly more planulae were shed by shallow water colonies 
than by deep water colonies (P < 0.0 1 ; Wilcoxon's signed rank test; Sokal and Rohlf, 
198 1 ). At the peak of the reproductive season about 20-80 planulae on average were 
shed per 100 g skeleton from shallow water colonies during one night. In deep water 
colonies the number did not exceed four planulae. An additional 55 deep water colo- 
nies were sampled during summer and fall (July-November) to examine whether the 
reproductive season there differs from that of shallow water populations. All histolog- 
ical sections were free of eggs. Moreover, no planulae were shed during a parallel 
study where branches were carefully broken in situ from an additional 80 colonies. 
These results indicate that the reproductive season of deep water colonies is probably 
two to three months shorter than that of shallow water populations. 

DISCUSSION 

The study of coral reproductive biology may be engaged with ambiguous defini- 
tions which could lead to wrong interpretations. For example, Fadlallah (1983) indi- 
cated that confusion arises from the applications of the term hermaphroditism, which 
describe two different life history processes: ( 1 ) development of monoecy over the 
lifetime of a specimen and, (2) sequential maturation of female and male products 
within one breeding period. Thus, it was accepted that 5". pistillata (Rinkevich and 
Loya, 1979a, b) and Goniastrea australensis (Kojis and Quinn, 198 la, b) were prot- 
androus hermaphrodites over their lifetime, but protogynous hermaphrodites in each 
single reproductive season. The present study indicates that either sexuality and/or 
fecundity may be completely altered from one reproductive season to the next. Her- 



342 B. RINKEVICH AND Y. LOYA 

nfaphroditic colonies which exhibited high fecundity in one season became male or 
even sterile thereafter, and vice versa. Small colonies (geometric mean radius 7 < 2 
cm) which invest much energy in rapid growth (Loya, 1985), possess only male go- 
nads ir their hrst year of reproduction. An increase in colony size correlated with an 
increa:,., in percentage of hermaphroditic colonies within the population (Rinkevich 
and Loya, 1979b). Reproduction of injured colonies of S. pistillata which invested 
energy in growth and regeneration was significantly reduced for at least two successive 
reproductive seasons after the fracturing event (Rinkevich, 1982). In addition, the 
fecundity of dying colonies was reduced several months before their death (Table III 
and Rinkevich and Loya, 1986), and dying colonies often changed their sexuality 
before their mortality and became male. Field experiments also demonstrated that 
the number of female gonads per polyp in S. pistillata was significantly reduced in 
colonies competing intraspecifically and the typical synchrony in reproduction 
among different branches of a given colony was changed and disynchronized (Rin- 
kevich and Loya, 1985). 

From the above results, we suggest that sexuality and fecundity in S. pistillata are 
responsive to the general state of health of the colony and its energetic limitations. 

Studies also addressed reproduction/energy allocation questions in other coral 
reef species. Kojis and Quinn (1985) found lower fecundity in damaged Goniastrea 
favulus colonies compared to unharmed controls and suggested that this resulted 
from reallocation of resources to growth activities that would repair damaged tissue 
and cover the broken skeletons. Richmond (1984) indicated that reef corals may 
allocate energy into new tissue via budding for colony growth, or via planulation 
for production of new colonies. He found that colonies of Pocillopora damicornis at 
Enewetak atoll, Marshall Islands, allocated the majority of their reproductive energy 
into larva production while in the eastern Pacific the same species channels energy 
into colony growth. Thus, internal and/or external (see below) determinants may 
play a significant role in the expression of sexuality or fecundity in hermatypic corals, 
although the mechanisms are not yet understood. 

Stimson (1978) proposed that coral species which release planulae are characteris- 
tic of shallow water environments such as reef flats, and hypothesized that shallow 
water species should planulate to facilitate early settlement in the parental habitat. 
Conversely, deep water corals should release eggs and sperm into the water to facili- 
tate dispersal. More recent studies, however, suggest that the mode of reproduction 
is related to more complex factors than habitat alone (Harriott, 1 983; Szmant, 1 986). 
Thus it is of interest to study the mode of reproduction of the same species in two 
different depths. Karlson (1981) found a reduction in reproductive activity with in- 
creasing depth in two Jamaican species ofZoanthus. Kojis and Quinn (1983) further 
indicate that fecundity of Acropora palifera decreased with depth. Colonies at depths 
greater than 1 2 m had approximately half the fecundity of surface colonies. These 
studies support the results of the present study (Tables IV, V, VI) which indicate high 
differences in fecundity between shallow and deep water 5". pistillata colonies. 

The importance of available energy for reproduction is apparent from the de- 
crease of fecundity in deep water populations. S. pistillata invests photosynthetically 
derived energy in reproduction (Rinkevich, 1982; Rinkevich and Loya, 1983). Mc- 
Closkey and Muscatine (1984) found that the daily CZAR (the percentage contribu- 
tion of zooxanthellae-translocated carbon to animal maintenance respiration) in S. 
pistillata in deep water was less than half of that in shallow water. Mean CZAR at 35 
m was 78%, compared to 157% at 3 m. They also found that the decreased carbon 
availability to the host animal at 35 m was the consequence of both decreased net 
carbon fixation and decreased percentage of net fixed carbon translocated to the host. 



VARIABILITY IN SEXUAL REPRODUCTION OF A CORAL 343 

Therefore, we suggest, that the generous daily carbon supply in shallow water colonies 
enables them to channel significantly more energy to reproduction than deep water 
colonies. 

It is hard t