CRC Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation

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Author: Leslie A. Dierauf | Frances M.D. Gulland

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CRC Handbook of

Marine Mammal Medicine Second Edition

0839_frame_FM1 Page 2 Wednesday, May 23, 2001 10:38 AM

Cover: In 1988, this marine mammal quilt was designed and constructed by scores of artists, needlework experts, and quilters to honor the efforts of The Marine Mammal Center (TMMC), in Sausalito, California. The quilt incorporates the designs of artists Richard Ellis, Pieter Folkens, Larry Foster, Dugald Stermer, and 25 others. The quilt travels on display, and to date has been exhibited at the California Academy of Sciences, the Monterey Bay Aquarium, and TMMC. This cover is in honor of the more than 800 volunteers who work at TMMC and for our contributors, reviewers, and editors. Thank you!

CRC Handbook of

Marine Mammal Medicine Second Edition Edited by

Leslie A. Dierauf and Frances M. D. Gulland

CRC Press Boca Raton London New York Washington, D.C.

0839_frame_FM1 Page 4 Tuesday, April 9, 2002 1:34 PM

Senior Editor: John Sulzycki Production Manager: Carol Whitehead Marketing Manager: Carolyn Spence

Illustrations in Chapters 9 and 19 are © Sentiel A. Rommel. Library of Congress Cataloging-in-Publication Data CRC Handbook of marine mammal medicine / edited by Leslie A. Dierauf and Frances M.D. Gulland.--2nd ed. p. cm. Includes bibliographical references and index. ISBN 0-8493-0839-9 (alk. paper) 1. Marine mammals--Diseases--Handbooks, manuals, etc. 2. Marine mammals--Health--Handbooks, manuals, etc. 3. Veterinary medicine--Handbooks, manuals etc. 4. Wildlife rehabilitation--Handbooks, manuals etc. I. Title: Handbook of marine mammal medicine. II. Dierauf, Leslie A., 1948- III. Gulland, Frances M. D. SP997.5.M35 C73 2001 636.9′5--dc21

2001025211 CIP

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the authors and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage or retrieval system, without prior permission in writing from the publisher. All rights reserved. Authorization to photocopy items for internal or personal use, or the personal or internal use of specific clients, may be granted by CRC Press LLC, provided that $1.50 per page photocopied is paid directly to Copyright clearance Center, 222 Rosewood Drive, Danvers, MA 01923 USA. The fee code for users of the Transactional Reporting Service is ISBN 0-8493-0839-9/01/$0.00+$1.50. The fee is subject to change without notice. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. The consent of CRC Press LLC does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from CRC Press LLC for such copying. Direct all inquiries to CRC Press LLC, 2000 N.W. Corporate Blvd., Boca Raton, Florida 33431. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe.

Visit the CRC Press Web site at www.crcpress.com © 2001 by CRC Press LLC No claim to original U.S. Government works International Standard Book Number 0-8493-0839-9 Library of Congress Card Number 2001025211 Printed in the United States of America 2 3 4 5 6 7 8 9 0 Printed on acid-free paper

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Dedication

This book is dedicated to Dr. Nancy Foster— A whole generation of veterinarians for whom you, as a scientist, were our mentor, our inspiration, and our motivation in our pursuit of marine science, policy, and marine mammal medicine, thank you. We miss you.

Thank you, Joe— for caring for Nancy for caring for the animals, and for being a leader for us in the field of marine mammal medicine.



Preface

Read not to contradict and confute, nor to believe and take for granted, nor to find talk and discourse, but to weigh and consider. —Francis Bacon, 1625

It has been more than 10 years since the first edition of the Handbook of Marine Mammal Medicine was published; during that time, the book has sold consistently (almost 2000 copies worldwide). Since its publication in 1990, there has been an exponential growth of experience and published literature addressing marine mammal medicine. Marine mammals have captured the imagination of not only the public, but also the scientific community. Despite this increase in information, much remains to be learned about the medicine of marine mammals. We hope that by sharing what is known to date, veterinarians will be encouraged to explore the unknown, and share this new information in the future. The meaning of the phrase “marine mammal medicine” has greatly expanded, and the contents of this second edition attempt to reflect this. As we enter the new millennium, veterinarians are not only involved in diagnosis and treatment of disease, but also in the bigger picture, including marine mammals as sentinels of ocean health, animal well-being, marine mammal strandings and unusual mortality events, legislation governing marine mammal health and population trends, and tagging and tracking of rehabilitated and released animals. To care for marine mammals effectively, veterinarians also need to understand their anatomy, physiology, and behavior. As the field develops, we must encourage new members of the profession and be able to advise students on careers in the field of marine mammal medicine. We hope our vision of what marine mammal medicine is in the 21st century becomes yours. With 66 contributors, and almost 100 reviewers, all working together to help craft 45 scientifically based chapters, we believe the contents of this textbook are light-years ahead of the topics presented in the first edition of the Handbook of Marine Mammal Medicine. For these extraordinary efforts, we wish to offer our utmost thanks to everyone involved. We appreciate the time taken away from their work to share their knowledge and experience with others. With all the reference books, journals, e-mails, and Web sites each author investigated, this second edition is an explosion of new information. We apologize for any current medical literature on marine mammals we may have inadvertently overlooked in this effort. Almost every year since 1995, CRC Press, the publisher of the first edition of the handbook, has contacted one of the editors (Dierauf ) asking if she “would be interested in publishing a second edition?” And almost every year since 1995, due to time constraints, more than full-time commitments elsewhere, and the fact that her current efforts are directed toward habitat protection for threatened and endangered species (U.S. Fish and Wildlife Service, Albuquerque, NM) and environmental education (co-founder and chair of the Alliance of Veterinarians for the Environment, Nashville, TN), she has emphatically and succinctly said “no.” Except, for early fall, 1999, when she hesitated . . . said she had to make a few phone calls, and would call back.


The phone calls were to the now coeditor (Gulland) and, although she too had been asked previously and declined, she too hesitated. It really was time, almost the 10-year anniversary of the first edition; there was so much new information, so many new scientists entering the field, and an amazing array of students calling for help in advancing future careers in marine mammal medicine. We agreed that it was indeed time, if we could persuade fellow colleagues to join us in this effort. To our great surprise and wonder, considering how pressed for time everyone is these days, more than 90% of the scientists we called enthusiastically agreed to participate. We called CRC in October 1999, and said, “yes.” We are proud of our authors and our publisher for bringing this second edition to publication promptly to ensure that the information presented is as up to date and future oriented as is possible in this age of information. The first edition of this book limited its scope to U.S. and Canadian issues and species. This edition tries harder to address international concerns and the worldwide practice of marine mammal medicine. We chose to write the text in (no, not English — sorry Frances!) American (phrases, spelling) for consistency with the first edition. Both metric and American measurements are provided, and there is a conversion table in the appendix. In the references at the end of each chapter, we include abstracts from conference proceedings (many of which can be found on the International Association for Aquatic Animal Medicine, or IAAAM, CD-ROM; see Chapters 7 and 8 for ordering information), as well as peer-reviewed books and journals. This is to provide the reader with as much current information as possible; the reader is encouraged to seek peer-reviewed journal articles by the same authors as their pieces are published. We have Web information from reputable sources within the context of each chapter (in bold), information from veterinary and marine scientists through personal communications (pers. comm.), unpublished data (unpubl. data), cross-referencing that refers to pertinent information in other chapters (see Chapter …), and an extensive index. The chapters in this second edition have been peer-reviewed. Yet, despite this peer-reviewed information, the editors still wish to emphasize that, in the practice of marine mammal medicine, nothing—not Web information, not journal information, not e-mail information—substitutes for talking to your peers and colleagues prior to performing a new procedure, or administering a pharmaceutical to a marine mammal. Nothing beats a healthy exchange of questions, answers, and experiences to assist in decision making. Again, we wish to thank everyone we have worked with over the past year (authors, coauthors, editors, peer-reviewers, colleagues) for giving us their unending support, for responding to our unceasing phone calls and e-mails, and for helping us maintain our enthusiasm. We thank Raymond Tarpley, David St. Aubin, Shannon Atkinson, and Bill Amos for wonderful lastminute rescues. We offer special thanks to the staff and volunteers at The Marine Mammal Center, in Sausalito, CA (we quietly refer to these Editorial and Literary Volunteers as our “elves”) for their consistent, constant, and voluntary efforts on behalf of this production. In particular, we thank Rebecca Duerr, Danielle Duggan, Denise Greig, Michelle Lander, Gayle Love, Alana Phillips, Kathryn Zagzebski, Kelly Alman, Amber Clutton-Brock, and Tanya Zabka. Thanks are due to Andy Draper for ensuring polar bears were not left out in the cold, and to both Andy Draper and Jim Hurley for keeping our spirits up. We could not have done this without the help of every one of you. Leslie A. Dierauf Frances M. D. Gulland


Editors

Leslie A. Dierauf, V.M.D, is a wildlife veterinarian and conservation biologist with 17 years of clinical veterinary practice experience, specializing in marine mammal and small animal emergency medicine. She currently works with the U.S. Fish and Wildlife Service (Service), primarily on habitat conservation planning efforts for all types of threatened and endangered species in Texas, Arizona, New Mexico, and Oklahoma. Her primary focus is forming partnerships between the federal government and the private sector/citizenry. Prior to joining the Service, she worked as a scientific advisor on committee staff for the U.S. House of Representatives in Washington, D.C. In 1998, Dr. Dierauf was honored by the profession of veterinary medicine with the American Veterinary Medical Association’s National Animal Welfare Award. She also served as an American Association for the Advancement of Science Congressional Science Fellow. Dr. Dierauf currently sits on the Marine Ecosystem Health Program Advisory Board, a research and science policy effort located on Orcas Island, WA, and associated with the University of California, Davis, Wildlife Health Center. She also served 8 years on the American Veterinary Medical Association’s Environmental Affairs Committee, and 8 years on the National Marine Fisheries Service’s Working Group on Marine Mammal Unusual Mortality Events. She is the co-founder and chair of the Board of the Alliance of Veterinarians for the Enviornment. Dr. Dierauf is a member of the International Association for Aquatic Animal Medicine, the Alliance of Veterinarians for the Environment, the Society for Conservation Biology, and the American Veterinary Medical Association. She lives in Santa Fe, NM, with Jim Hurley, her partner of 22 years, and their three dogs. Frances M. D. Gulland, Vet. M.B., M.R.C.V.S., Ph.D., is a veterinarian interested in the role of disease in wildlife conservation. She obtained her veterinary degree from the University of Cambridge (England) in 1984 and her Ph.D., also from the University of Cambridge (Zoology Department) in 1991. Dr. Gulland worked at the Zoological Society of London as House Surgeon and later as Fellow in Wildlife Diseases, before moving to California in 1994. Dr. Gulland was introduced to marine mammals by her father, John A. Gulland, but became involved in their medicine when she started to work at The Marine Mammal Center (TMMC), Sausalito, CA, in 1994. As Director of Veterinary Services at TMMC, Dr. Gulland is involved in marine mammal strandings, rehabilitation, and disease investigation. She learns about marine mammal medicine on a daily basis from the animals and people around her. Dr. Gulland currently serves as a scientific advisor to the Oiled Wildlife Care Network in California and the Marine Mammal Commission, and is a member of the Working Group on Marine Mammal Unusual Mortality Events, the International Association for Aquatic Animal Medicine, the Wildlife Disease Association, and the Society for Marine Mammalogy.



Contributors

Brian M. Aldridge B.V.Sc., Ph.D., A.C.V.I.M. Department of Pathology, Microbiology, and Immunology School of Veterinary Medicine University of California Davis, California William Amos, Ph.D. Department of Zoology University of Cambridge Cambridge, England Brad F. Andrews SeaWorld of Florida Orlando, Florida Jim Antrim SeaWorld of California San Diego, California Kristen D. Arkush, Ph.D. Bodega Marine Laboratory University of California Bodega Bay, California Shannon K. C. Atkinson, Ph.D. Alaska SeaLife Center and University of Alaska Seward, Alaska Cathy A. Beck, M.S. U.S. Geological Survey Florida Caribbean Science Center Sirenia Project Gainesville, Florida Robert K. Bonde, Ph.D. U.S. Geological Survey Florida Caribbean Science Center Sirenia Project Gainesville, Florida

Gregory D. Bossart, V.M.D., Ph.D. Division of Marine Mammal Research and Conservation Harbor Branch Oceanographic Institution Fort Pierce, Florida Michael Brent Briggs, D.V.M. Brookfield Zoo Brookfield, Illinois Fiona Brook, Ph.D., R.D.M.S., D.C.R. Department of Optometry and Radiography The Hong Kong Polytechnic University Hung Hom, Kowloon, Hong Kong John D. Buck, Ph.D. Mote Marine Laboratory Sarasota, Florida Daniel F. Cowan, M.D. Department of Pathology University of Texas Medical Branch Galveston, Texas Murray D. Dailey, Ph.D. The Marine Mammal Center Marin Headlands Sausalito, California Leslie M. Dalton, D.V.M. SeaWorld of Texas San Antonio, Texas Leslie A. Dierauf, V.M.D. Alliance of Veterinarians for the Environment Santa Fe, New Mexico Samuel R. Dover, D.V.M. Santa Barbara Zoological Garden Santa Barbara, California


Deborah A. Duffield, Ph.D. Department of Biology Portland State University Portland, Oregon J. Lawrence Dunn, V.M.D. Department of Research and Veterinary Medicine Mystic Aquarium Mystic, Connecticut Ruth Y. Ewing, D.V.M. National Marine Fisheries Service South East Florida Science Center Miami, Florida Salvatore Frasca, Jr., V.M.D., Ph.D. Department of Pathobiology University of Connecticut Storrs, Connecticut Laurie J. Gage, D.V.M. Six Flags MarineWorld Vallejo, California Edward V. Gaynor, D.V.M. SeaWorld of Florida Orlando, Florida Scott Gearhart, D.V.M. SeaWorld of Florida Orlando, Florida Leah L. Greer, D.V.M. Department of Comparative Medicine College of Veterinary Medicine University of Tennessee Knoxville, Tennessee Frances M. D. Gulland, Vet. M.B., M.R.C.V.S., Ph.D. The Marine Mammal Center Marin Headlands Sausalito, California Martin Haulena, M.Sc., D.V.M. The Marine Mammal Center Marin Headlands Sausalito, California

Robert Bruce Heath, D.V.M., M.Sc., Dipl. A.C.V.A. Fort Collins, Colorado Aleta A. Hohn National Marine Fisheries Service Beaufort Laboratory Beaufort, North Carolina Carol House, Ph.D. Cutchogue, New York James A. House, D.V.M., Ph.D. Cutchogue, New York Eric D. Jensen, D.V.M. U.S. Navy Marine Mammal Program San Diego, California Suzanne Kennedy-Stoskopf, D.V.M., Ph.D., Dipl. A.C.Z.M. North Carolina State University Raleigh, North Carolina Donald P. King, Ph.D. Department of Pathology, Microbiology and Immunology School of Veterinary Medicine University of California Davis, California Michelle E. Lander, M.Sc. The Marine Mammal Center Marin Headlands Sausalito, California Lynn W. Lefebvre, Ph.D. U.S. Geological Survey Florida Caribbean Science Center Sirenia Project Gainesville, Florida Linda J. Lowenstine, D.V.M., Ph.D., Dipl. A.C.V.P. Department of Pathology, Microbiology, and Immunology School of Veterinary Medicine University of California Davis, California


James F. McBain, D.V.M. SeaWorld of California San Diego, California Ted Y. Mashima, D.V.M., Dipl. A.C.Z.M. Center for Government and Corporate Veterinary Medicine University of Maryland Baltimore, Maryland Debra Lee Miller, D.V.M., Ph.D. Division of Comparative Pathology University of Miami School of Medicine Miami, Florida Michael J. Murray, D.V.M. Monterey Bay Aquarium Monterey, California Daniel K. Odell, Ph.D. SeaWorld of Florida Orlando, Florida Todd M. O’Hara, D.V.M., Ph.D. North Slope Borough Department of Wildlife Management Barrow, Alaska Thomas J. O’Shea, M.S., Ph.D. U.S. Geological Survey Midcontinent Ecological Science Center Fort Collins, Colorado Michelle Lynn Reddy SAIC Maritime Services San Diego, California

Sentiel A. Rommel, Ph.D. Eckerd College Florida Marine Research Institute Marine Mammal Pathobiology Laboratory St. Petersburg, Florida Teri K. Rowles, D.V.M., Ph.D. Office of Protected Resources National Marine Fisheries Service Silver Spring, Maryland David J. St. Aubin, Ph.D. Mystic Aquarium Mystic, Connecticut Sara L. Shapiro Florida Fish and Wildlife Conservation Commission Florida Marine Research Institute St. Petersburg, Florida Terry R. Spraker, D.V.M., Ph.D., Dipl. A.C.V.P. Diagnostic Laboratory College of Veterinary Medicine Colorado State University Fort Collins, Colorado Michael K. Stoskopf, D.V.M., Ph.D., Dipl. A.C.Z.M. Environmental Medicine Consortium College of Veterinary Medicine North Carolina State University Raleigh, North Carolina

Thomas H. Reidarson, D.V.M., Dipl. A.C.Z.M. SeaWorld of California San Diego, California

Jeffrey L. Stott, Ph.D. Department of Pathology, Microbiology, and Immunology School of Veterinary Medicine University of California Davis, California

Michael G. Rinaldi, D.V.M. Department of Pathology University of Texas Health Science Center San Antonio, Texas

Jay C. Sweeney, V.M.D. Dolphin Quest San Diego, California

Todd R. Robeck, D.V.M., Ph.D SeaWorld of Texas San Antonio, Texas

Forrest I. Townsend, Jr., D.V.M. Bayside Hospital for Animals Fort Walton Beach, Florida


Pamela Tuomi, D.V.M. Alaska SeaLife Center Seward, Alaska William Van Bonn, D.V.M. U.S. Navy Marine Mammal Program San Diego, California Frances M. Van Dolah, Ph.D. National Ocean Services Charleston, South Carolina Michael T. Walsh, D.V.M. SeaWorld of Florida Orlando, Florida Andrew J. Westgate, Ph.D. Duke Marine Laboratory Beaufort, North Carolina

Janet Whaley, D.V.M. Office of Protected Resources National Marine Fisheries Service Silver Spring, Maryland Scott Willens, D.V.M. North Carolina State University Raleigh, North Carolina Graham A. J. Worthy, Ph.D. Department of Biology University of Central Florida Orlando, Florida Nina M. Young, M.S. Center for Marine Conservation Washington, D.C.


Contents

Section I 1

Emerging Pathways in Marine Mammal Medicine

Marine Mammals as Sentinels of Ocean Health Michelle Lynn Reddy, Leslie A. Dierauf, and Frances M. D. Gulland

Introduction ................................................................................................3 Sentinels......................................................................................................3 Ecosystem Changes Detected by Sentinels..............................................4 Marine Mammals as Sentinels..................................................................5 Conclusion ..................................................................................................9 Acknowledgments ......................................................................................9 References ...................................................................................................9

2

Emerging and Resurging Diseases Debra Lee Miller, Ruth Y. Ewing, and Gregory D. Bossart

Introduction ..............................................................................................15 Cetaceans ..................................................................................................16 Pinnipeds...................................................................................................19 Manatees ...................................................................................................22 Sea Otters..................................................................................................23 Polar Bears ................................................................................................24 Conclusion ................................................................................................24 Acknowledgments ....................................................................................25 References .................................................................................................25

3

Florida Manatees: Perspectives on Populations, Pain, and Protection Thomas J. O’Shea, Lynn W. Lefebvre, and Cathy A. Beck

Introduction ..............................................................................................31 Maiming of Manatees in Collisions with Boats ....................................33 A Primer on Manatee Population Biology: Accounting for the Confusion and Uncertainty.....................................................36 Estimation of Population Size and Trend.....................................36 Carcass Counts, Mortality, and Survival......................................39


Population Models..........................................................................40 Uncertainties on Population Status: A Red Herring? ...........................40 References .................................................................................................42

4

Marine Mammal Stranding Networks Frances M. D. Gulland, Leslie A. Dierauf, and Teri K. Rowles

Introduction ..............................................................................................45 Objectives of Stranding Networks ..........................................................45 Stranding Networks Worldwide ..............................................................46 Acknowledgments ....................................................................................66 References .................................................................................................66

5

Marine Mammal Unusual Mortality Events Leslie A. Dierauf and Frances M. D. Gulland

Introduction ..............................................................................................69 MMUME Responses in the United States .............................................70 The U.S. National Contingency Plan ...........................................71 Expert Working Group on MMUMEs...........................................71 The MMUME Response.................................................................74 MMUME Fund................................................................................76 Lessons Learned........................................................................................77 The Cooperative Response ............................................................77 The Process .....................................................................................78 UMMME Fund................................................................................78 Results Accrued from Title IV of the MMPA........................................78 How Can You Help? ................................................................................79 Conclusion ................................................................................................79 Acknowledgments ....................................................................................79 References .................................................................................................79

6

Mass Strandings of Cetaceans Michael T. Walsh, Ruth Y. Ewing, Daniel K. Odell, and Gregory D. Bossart

Introduction ..............................................................................................83 Theories to Explain Mass Strandings .....................................................83 Current Investigations into Mass Strandings ........................................86 Evaluation of a Mass Stranding ..............................................................87 Management of a Mass Stranding...........................................................88 Disposition of Animals in a Mass Stranding .........................................92 Euthanasia .......................................................................................94 Return to the Sea ...........................................................................94 Survival of Treated Whales............................................................94


Conclusion ................................................................................................94 Acknowledgments ....................................................................................95 References .................................................................................................95

7

Careers in Marine Mammal Medicine Leslie A. Dierauf, Salvatore Frasca, Jr., and Ted Y. Mashima

Introduction ..............................................................................................97 Full-Time Employment..................................................................97 Part-Time Employment..................................................................98 Personality Traits and Other Tools...............................................98 Summary .........................................................................................99 The Six-Step Method for Landing That Perfect JobWorking with Marine Mammals ........................................................................99 1. The First Step—Taking a Personal Self-Assessment ...............99 2. The Second Step—Categorizing Your Unique Skills, Strategies, and Approaches ......................................................100 3. The Third Step—Planning for Action and Timing................102 4. The Fourth Step—Making Choices ........................................102 5. The Fifth Step—Preparing for the Interview..........................103 6. The Sixth Step—Starting Your New Job ................................106 Accessing Resources ..............................................................................107 Internships and Residencies ........................................................107 Matched Internships.........................................................107 Matched Residencies ........................................................108 Other Internships..............................................................108 Graduate Degree Programs ..........................................................109 Other Related Programs...............................................................110 Advanced Training Programs.......................................................111 Fellowships ...................................................................................112 Scientific Societies and Membership Organizations .................112 Recommendations and Conclusions.....................................................113 Acknowledgments ..................................................................................114 References ...............................................................................................114

8

The Electronic Whale Leslie A. Dierauf

Introduction ............................................................................................117 Using Your Head on the Web................................................................117 Reference Databases...............................................................................118 General Biomedical and Veterinary Medical Sites ....................118 Model Web Sites and Evidence-Based Medicine ........................119


Marine Mammal–Related Listserves...........................................120 Other Internet Discussion and Marine Mammal Information Lists ......................................................................121 Online Marine Mammal Journals and Textbooks .....................121 Fellowships, Foundations, and Grants........................................122 Fellowships........................................................................122 Foundations .......................................................................123 Grants ................................................................................123 Federal Government Listings ......................................................123 Miscellaneous Electronic Resources ...........................................123 Meetings and Proceedings on CD-ROM.....................................125 Electronic Addresses for Other Chapters in This Book ............125 Disclaimer...............................................................................................126 Conclusions ............................................................................................126 References ...............................................................................................126

Section II 9

Anatomy and Physiology of Marine Mammals

Gross and Microscopic Anatomy Sentiel A. Rommel and Linda J. Lowenstine

Introduction ............................................................................................129 External Features....................................................................................138 Sea Lions .......................................................................................138 Manatees .......................................................................................138 Seals...............................................................................................139 Dolphins ........................................................................................139 Microanatomy of the Integument.........................................................139 The Superficial Skeletal Muscles..........................................................141 The Diaphragm as a Separator of the Body Cavities ..........................142 Gross Anatomy of Structures Cranial to the Diaphragm ...................142 Heart and Pericardium .................................................................142 Pleura and Lungs ..........................................................................143 Mediastinum .................................................................................143 Thymus .........................................................................................143 Thyroids ........................................................................................143 Parathyroids ..................................................................................144 Larynx............................................................................................144 Caval Sphincter ............................................................................144 Microscopic Anatomy of Structures Cranial to the Diaphragm ........144 Respiratory System.......................................................................144 Thymus .........................................................................................145 Thyroids ........................................................................................145 Parathyroids ..................................................................................145


Gross Anatomy of Structures Caudal to the Diaphragm ...................145 Liver...............................................................................................145 Digestive System ..........................................................................145 Urinary Tract ................................................................................147 Genital Tract.................................................................................147 Adrenal Glands .............................................................................148 Microscopic Anatomy of Structures Caudal to the Diaphragm.........148 Liver...............................................................................................148 Digestive System ..........................................................................148 Urinary Tract ................................................................................149 Genital Tract.................................................................................149 Adrenals ........................................................................................150 Lymphoid and Hematopoietic Systems ......................................150 Nervous System .....................................................................................150 Circulatory Structures ...........................................................................151 The Potential for Thermal Insult to Reproductive Organs ................152 Skeleton ..................................................................................................153 Ribs................................................................................................155 Sternum.........................................................................................155 Postthoracic Vertebrae .................................................................156 Sacral Vertebrae ............................................................................156 Chevron Bones..............................................................................156 Pectoral Limb Complex ...............................................................156 Pelvic Limb Complex...................................................................157 Sexual Dimorphisms ....................................................................157 Bone Marrow.................................................................................158 Acknowledgments ..................................................................................158 References ...............................................................................................158

10 Endocrinology David J. St. Aubin

Introduction ............................................................................................165 Sample Collection and Handling ..........................................................166 Blood..............................................................................................166 Saliva .............................................................................................166 Feces ..............................................................................................166 Urine..............................................................................................166 Tissues...........................................................................................167 Pineal Gland ...........................................................................................167 Hypothalamus–Pituitary........................................................................169 Thyroid Gland ........................................................................................169 Adrenal Gland ........................................................................................177


Osmoregulatory Hormones ...................................................................182 Vasopressin....................................................................................183 Renin–Angiotensin System..........................................................185 Atrial Natriuretic Peptide............................................................185 Endocrine Pancreas ................................................................................185 Future Studies.........................................................................................186 Acknowledgments ..................................................................................187 References ...............................................................................................187

11 Reproduction Todd R. Robeck, Shannon K. C. Atkinson, and Fiona Brook

Introduction ............................................................................................193 Physiology of Reproduction...................................................................193 Pinniped Reproduction ..........................................................................195 Female Pinniped Reproduction ...................................................195 Reproductive Cycle ..........................................................195 Estrous Cycle ....................................................................196 Pregnancy and Pseudopregnancy .....................................197 Embryonic Diapause and Reactivation ...........................198 Implantation......................................................................198 Pregnancy Diagnosis.........................................................199 Induction of Parturition ...................................................199 Lactation............................................................................200 Milk Collection ................................................................200 Male Pinniped Reproduction .......................................................200 Anatomy ............................................................................200 Sexual Maturity ................................................................201 Seasonality ........................................................................201 Contraception and Control of Aggression ..................................202 Females ..............................................................................202 Males .................................................................................202 Reproductive Abnormalities in Pinnipeds .................................203 Cetacean Reproduction..........................................................................204 Female Cetacean Reproduction...................................................204 Reproductive Maturity .....................................................204 Bottlenose Dolphin ...........................................204 White-Sided Dolphin ........................................204 Killer Whale.......................................................204 False Killer Whale .............................................205 Beluga.................................................................205 Reproductive Cycle ..........................................................205 Bottlenose Dolphin ...........................................205 White-Sided Dolphin ........................................205


Killer Whale.......................................................206 False Killer Whale .............................................206 Beluga.................................................................206 Estrous Cycle and Ovarian Physiology ...........................206 Bottlenose Dolphin ...........................................206 Killer Whale.......................................................208 False Killer Whale .............................................208 Suckling (Lactational) Suppression of Estrus .................209 Corpora Albicantia and Asymmetry of Ovulation ........210 Pseudopregnancy...............................................................210 Pregnancy ..........................................................................211 Bottlenose Dolphin ...........................................211 Killer Whale.......................................................211 Beluga.................................................................212 Pregnancy Diagnosis.........................................................212 Parturition .........................................................................212 Stages of Parturition .........................................212 Induction of Parturition ...................................212 Male Cetacean Reproduction ......................................................215 Sexual Maturity ................................................................215 Bottlenose Dolphin ...........................................215 White-Sided Dolphin ........................................215 Killer Whale.......................................................215 Beluga.................................................................216 Seasonality ........................................................................216 Bottlenose Dolphin ...........................................216 White-Sided Dolphin ........................................216 Killer Whale.......................................................217 False Killer Whale .............................................217 Beluga.................................................................217 Contraception and Control of Aggression ..................................217 Females ..............................................................................217 Males .................................................................................218 Reproductive Abnormalities in Cetaceans.................................218 Artificial Insemination.................................................................219 Semen Collection and Storage.........................................219 Manipulation and Control of Ovulation.........................221 Induction of Ovulation .....................................221 Synchronization of Ovulation..........................222 Insemination Techniques .................................................223 Future Applications ..........................................................224 Acknowledgments ..................................................................................225 References ...............................................................................................226

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12 Immunology Donald P. King, Brian M. Aldridge, Suzanne Kennedy-Stoskopf, and Jeffrey L. Stott

Introduction ............................................................................................237 Overview of the Immune System.........................................................238 Innate Immunity and the Inflammatory Response ...................238 Adaptive Immune Response ........................................................238 Cytokines ......................................................................................239 Immunodiagnostics ................................................................................240 Inflammation ................................................................................240 Cellular Immunity .......................................................................241 Functional Immune Testing ........................................................242 In Vitro ..............................................................................242 In Vivo ...............................................................................242 Humoral Immunity ......................................................................243 Measurement of Pathogen-Specific Antibodies (Serodiagnostics) ......243 Serum/Virus Neutralization Test ................................................244 Precipitation/Agglutination Techniques.....................................244 Enzyme-Linked Immunosorbent Assay ......................................245 Total Immunoglobulin .................................................................245 Clinical Approach to Suspected Marine Mammal Immunological Disorders...................................................................246 Conclusion ..............................................................................................248 Acknowledgments ..................................................................................248 References ...............................................................................................248

13 Stress and Marine Mammals David J. St. Aubin and Leslie A. Dierauf

Introduction ............................................................................................253 Stressors ..................................................................................................253 Stress Response and Regulation............................................................254 Neurological Factors ....................................................................255 Endocrine Factors .........................................................................256 Catecholamines.................................................................256 Glucocorticoids.................................................................256 Mineralocorticoids............................................................260 Thyroid Hormones ...........................................................260 Other Hormones ...............................................................261 Immunological Factors.................................................................261 Indicators of Acute and Chronic Stress................................................262 Acute Response ............................................................................262 Chronic Response.........................................................................263 Future Research......................................................................................264


Conclusion ..............................................................................................265 Acknowledgments ..................................................................................265 References ...............................................................................................265

14 Genetic Analyses Deborah A. Duffield and William Amos

Introduction ............................................................................................271 Genetic Techniques ...............................................................................271 DNA Sequencing ..........................................................................271 “Tandem Repeats” and DNA Fingerprinting .............................272 Genetic Analyses Applied to Stranded Marine Mammals..................272 Species Identification ...................................................................273 Population Identification .............................................................273 Social Organization ......................................................................274 Genetic Analysis Applied to Captive Maintenance and Breeding Programs ..............................................................................275 Paternity Testing ..........................................................................275 Hybrid Detection..........................................................................276 Sampling .................................................................................................277 Conclusion ..............................................................................................278 Acknowledgments ..................................................................................278 References ...............................................................................................278

Section III

Infectious Diseases of Marine Mammals

15 Viral Diseases Suzanne Kennedy-Stoskopf

Introduction ............................................................................................285 Virus Isolation—An Overview ..............................................................285 Poxviruses ...............................................................................................286 Host Range....................................................................................286 Clinical Signs................................................................................287 Therapy .........................................................................................287 Pathology.......................................................................................287 Diagnosis .......................................................................................288 Differentials ..................................................................................288 Epidemiology ................................................................................289 Public Health Significance...........................................................289 Papillomaviruses ....................................................................................289 Host Range....................................................................................289 Clinical Signs................................................................................290


Therapy .........................................................................................290 Pathology.......................................................................................290 Diagnosis .......................................................................................290 Differentials ..................................................................................290 Epidemiology ................................................................................291 Public Health Significance...........................................................291 Adenoviruses ..........................................................................................291 Host Range....................................................................................291 Clinical Signs................................................................................291 Therapy .........................................................................................291 Pathology.......................................................................................292 Diagnosis .......................................................................................292 Epidemiology ................................................................................292 Public Health Significance...........................................................292 Herpesviruses..........................................................................................292 Host Range....................................................................................292 Virology .........................................................................................293 Clinical Signs................................................................................293 Therapy .........................................................................................294 Pathology.......................................................................................294 Diagnosis .......................................................................................294 Differentials ..................................................................................295 Epidemiology ................................................................................295 Public Health Significance...........................................................295 Morbilliviruses .......................................................................................296 Host Range....................................................................................296 Virology .........................................................................................296 Clinical Signs................................................................................296 Therapy .........................................................................................297 Pathology.......................................................................................297 Diagnosis .......................................................................................297 Differentials ..................................................................................297 Epidemiology ................................................................................298 Public Health Significance...........................................................298 Influenza Viruses ....................................................................................298 Host Range....................................................................................298 Clinical Signs................................................................................298 Therapy .........................................................................................299 Pathology.......................................................................................299 Diagnosis .......................................................................................299 Differentials ..................................................................................299 Epidemiology ................................................................................299 Public Health Significance...........................................................300


Caliciviruses (San Miguel Sea Lion Virus) ...........................................300 Host Range....................................................................................300 Clinical Signs................................................................................300 Therapy .........................................................................................300 Pathology.......................................................................................301 Diagnosis .......................................................................................301 Epidemiology ................................................................................301 Public Health Significance...........................................................302 Other Viruses..........................................................................................302 Hepadnavirus ................................................................................302 Coronavirus...................................................................................302 Retrovirus......................................................................................302 Rhabdoviruses...............................................................................303 Acknowledgments ..................................................................................303 References ...............................................................................................303

16 Bacterial Diseases of Cetaceans and Pinnipeds J. Lawrence Dunn, John D. Buck, and Todd R. Robeck

Introduction ............................................................................................309 Microbial Sampling Techniques............................................................310 Specific Bacterial Diseases of Cetaceans and Pinnipeds .....................312 Septicemia.....................................................................................312 Brucellosis .....................................................................................312 Cetaceans ..........................................................................313 Pinnipeds ...........................................................................314 Vibriosis.........................................................................................314 Cetaceans ..........................................................................315 Pinnipeds ...........................................................................315 Pasteurellosis ................................................................................315 Cetaceans ..........................................................................315 Pinnipeds ...........................................................................315 Erysipelothrix................................................................................316 Cetaceans ..........................................................................316 Pinnipeds ...........................................................................318 Mycobacterial Disease .................................................................319 Cetaceans ..........................................................................319 Pinnipeds ...........................................................................319 Leptospirosis .................................................................................320 Pinnipeds ...........................................................................320 Nocardia ........................................................................................321 Cetaceans ..........................................................................322 Pinnipeds ...........................................................................325


Miscellaneous Bacterial Disease ...........................................................325 Respiratory Disease ......................................................................325 Dermatological Disease ...............................................................326 Urogenital Disease .......................................................................327 Gastrointestinal Disease ..............................................................327 Conclusion ..............................................................................................328 Acknowledgments ..................................................................................328 References ...............................................................................................328

17 Mycotic Diseases Thomas H. Reidarson, James F. McBain, Leslie M. Dalton, and Michael G. Rinaldi

Introduction ............................................................................................337 Mycotic Diseases....................................................................................337 Epidemiology of Fungi ...........................................................................338 Modes of Transmission ................................................................338 Mechanisms of Pathogenesis.......................................................338 Clinical Manifestations .........................................................................339 Clinical Diagnostic Features of the Fungi ...........................................340 Therapeutics ...........................................................................................349 Conclusion ..............................................................................................351 Acknowledgments ..................................................................................352 References ...............................................................................................352

18 Parasitic Diseases Murray D. Dailey

Introduction ............................................................................................357 Removal and Fixation of Parasites for Identification..........................357 Treatment ...............................................................................................359 Parasites of Cetacea ...............................................................................359 Protozoa.........................................................................................359 Ciliates ..............................................................................359 Apicomplexans..................................................................359 Flagellates ..........................................................................360 Sarcodina ...........................................................................360 Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans)....................................................................361 Gastrointestinal Tract ......................................................361 Liver ...................................................................................365 Respiratory System, Sinuses, and Brain..........................365 Urogenital System ............................................................366 Connective Tissue ............................................................366


Ectoparasites .................................................................................367 Parasites of Pinnipeds ............................................................................367 Protozoa.........................................................................................367 Apicomplexans..................................................................367 Flagellates ..........................................................................368 Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans)....................................................................369 Gastrointestinal Tract ......................................................369 Respiratory and Circulatory Systems .............................370 Liver, Biliary System, and Pancreas ................................372 Connective Tissue ............................................................372 Ectoparasites .................................................................................372 Parasites of Sirenia .................................................................................372 Protozoa—Apicomplexans ...........................................................372 Helminths (Nematodes, Trematodes) .........................................373 Parasites of Sea Otters ...........................................................................373 Protozoa—Apicomplexans ...........................................................373 Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans)....................................................................373 Parasites of Polar Bears ..........................................................................374 Acknowledgments ..................................................................................374 References ...............................................................................................374

Section IV

Pathology of Marine Mammals

19 Clinical Pathology Gregory D. Bossart, Thomas H. Reidarson, Leslie A. Dierauf, and Deborah A. Duffield

Introduction ............................................................................................383 Abnormalities and Artifacts..................................................................383 Blood Collection.....................................................................................384 Sampling Equipment and Processing ..........................................384 Blood Collection Sites..................................................................385 Cetaceans ..........................................................................385 Otariids ..............................................................................385 Phocids ..............................................................................385 Odobenids..........................................................................385 Manatees ...........................................................................387 Sea Otters ..........................................................................387 Polar Bears.........................................................................390 Hematology (CBC)..................................................................................390 Evaluation of Erythrocytes ....................................................................391 Indices ...........................................................................................391


Anemia ..........................................................................................399 Classification of Anemia by RBC Indices ......................400 Normocytic, Normochromic ...........................400 Macrocytic, Hypochromic ................................400 Macrocytic, Normochromic .............................400 Microcytic, Normochromic, or Hypochromic ............................................401 Evaluation of Leukocytes ......................................................................401 Neutrophils or Heterophils..........................................................401 Eosinophils ....................................................................................401 Basophils .......................................................................................402 Monocytes and Lymphocytes ......................................................402 Leukocytes and Age .....................................................................402 Leukocytes and Disease ...............................................................403 Serum Analytes and Enzymes...............................................................403 Glucose, Lipids, and Pancreatic Enzymes ..................................403 Total Cholesterol and Triglycerides............................................404 Amylase, Lipase, and Trypsin-Like Immunoreactivity .............405 Markers of Hepatobiliary System Disorders ........................................406 Alanine Aminotransferase (ALT or SGPT) .................................406 Aspartate Aminotransferase (AST or SGOT) .............................407 Sorbitol Dehydrogenase (SDH) and Glutamate Dehydrogenase (GLDH) ...........................................................407 Lactate Dehydrogenase (LDH) .....................................................408 -Glutamyltransferase (GGT)......................................................408 Alkaline Phosphatase (ALP).........................................................409 Bilirubin ........................................................................................410 Bile Acids ......................................................................................411 Kidney-Associated Serum Analytes ......................................................411 Urea Nitrogen and Creatinine.....................................................411 Serum Proteins .......................................................................................413 Hematocrit and Total Plasma Protein ........................................413 Albumins and Globulins..............................................................414 Electrolytes .............................................................................................416 Sodium ..........................................................................................416 Potassium ......................................................................................416 Chloride.........................................................................................417 Total Carbon Dioxide...................................................................417 Calcium, Phosphorus, and Magnesium ......................................418 Calcium .............................................................................418 Phosphorus .......................................................................419 Magnesium ........................................................................419 Miscellaneous Serum Analytes .............................................................420 Uric Acid.......................................................................................420 Creatinine Phosphokinase ...........................................................420


Hemostatic Parameters..........................................................................420 Blood Types ...................................................................................420 Screening for Hemostatic Disorders ...........................................420 Prothrombin Time and Partial Prothrombin Time ...................421 Markers of Inflammation.......................................................................422 Erythrocyte Sedimentation Rate .................................................422 Serum Iron ....................................................................................422 Bone Marrow Evaluation .......................................................................423 Urinalysis ................................................................................................423 Conclusion ..............................................................................................424 Clinical Cases.........................................................................................424 Cetaceans ......................................................................................424 CASE 1—Bottlenose Dolphin ............................................424 History ...............................................................424 Clinicopathological Findings............................424 Discussion .........................................................424 CASE 2—Bottlenose Dolphin ............................................424 History ...............................................................424 Clinicopathological Findings............................424 Treatment ..........................................................424 Progress ..............................................................424 Additional Clinicopathological Findings.........425 Further Treatment.............................................425 CASE 3—Bottlenose Dolphin ............................................425 History ...............................................................425 Clinicopathological Findings............................425 Treatment ..........................................................425 Discussion .........................................................425 CASE 4—Killer Whale........................................................425 History ...............................................................425 Diagnosis ...........................................................425 Treatment ..........................................................425 Discussion .........................................................426 CASE 5—Killer Whale........................................................426 History ...............................................................426 Clinicopathological Findings............................426 Discussion .........................................................426 CASE 6—Pacific White-Sided Dolphin .............................426 History ...............................................................426 Clinicopathological Findings............................426 Treatment ..........................................................426 Subsequent Clinicopathological Findings .......427 Additional Treatment .......................................427


Diagnosis ...........................................................427 Discussion .........................................................427 Pinnipeds .......................................................................................427 CASE 1—Harbor Seal .........................................................427 History ...............................................................427 Clinicopathological Findings............................427 Treatment ..........................................................428 Post-Mortem Diagnosis ....................................428 Discussion .........................................................428 Manatees .......................................................................................428 CASE 1.................................................................................428 History ...............................................................428 Clinicopathological Findings............................428 Diagnosis ...........................................................428 Treatment ..........................................................428 Discussion .........................................................428 Sea Otters......................................................................................429 CASE 1.................................................................................429 History ...............................................................429 Clinicopathological Data..................................429 Radiographic Results ........................................429 Treatment ..........................................................429 Further Clinicopathological Data ....................429 Treatment ..........................................................429 Clinicopathological Data..................................429 Histopathological Diagnosis.............................429 Acknowledgments ..................................................................................430 References ...............................................................................................430

20 Cetacean Cytology Jay C. Sweeney and Michelle Lynn Reddy

Introduction ............................................................................................437 Sample Collection ..................................................................................438 Collection of Respiratory Tract Samples....................................438 Collection of Gastric Samples.....................................................438 Collection of Fecal Samples ........................................................439 Collection of Urinary Tract Samples..........................................439 Collection of Aspirates from Masses ..........................................439 Slide Preparation ....................................................................................439 Examination of Specimens ....................................................................441 Determination of Cellular Concentration within Slide Preparation.......................................................................441


Mucus............................................................................................441 Amorphous Material ....................................................................441 Interpretation ..........................................................................................441 Color..............................................................................................441 Epithelial Cells .............................................................................441 Leukocytes ....................................................................................442 Erythrocytes ..................................................................................442 Respiratory Tract....................................................................................442 Normal Findings...........................................................................442 Significant Findings......................................................................443 Stomach ..................................................................................................444 Normal Findings...........................................................................444 Significant Findings......................................................................444 Colon/Rectum ........................................................................................445 Normal Findings...........................................................................445 Significant Findings......................................................................445 Urinary Tract ..........................................................................................445 Normal Findings...........................................................................445 Significant Findings......................................................................446 Acknowledgments ..................................................................................446 References ...............................................................................................446

21 Gross Necropsy and Specimen Collection Protocols Teri K. Rowles, Frances M. Van Dolah, and Aleta A. Hohn

Introduction ............................................................................................449 Necropsy Examinations and Specimen Collection .............................450 Carcass Condition Code ........................................................................453 Morphometrics .......................................................................................453 Morphometric Data Protocol...........................................453 Genetics ..................................................................................................453 Genetic Sample Protocol..................................................454 Stomach Contents..................................................................................454 Stomach Contents Protocol .............................................454 Age...........................................................................................................454 Age Protocol......................................................................456 Reproductive Status ...............................................................................456 Reproductive Status Protocol ..........................................457 Pathology—Gross Necropsy Examination............................................457 Human Interactions .....................................................................458 Histopathology .......................................................................................458 Histopathology Protocol...................................................459 Acoustic Pathology ................................................................................459 Acoustic Pathology Protocol............................................460


Infectious Diseases.................................................................................460 Bacteriology...................................................................................460 Bacteriology Protocol........................................................460 Virology ...................................................................................................462 Virology Protocol ..............................................................462 Parasitology.............................................................................................462 Parasitology Protocol........................................................462 Non-Infectious Diseases ........................................................................464 Toxicology .....................................................................................464 Toxicology Protocol ..........................................................464 Harmful Algal Blooms ...........................................................................465 Harmful Algal Bloom Protocol ........................................467 Conclusions ............................................................................................467 Acknowledgments ..................................................................................467 References ...............................................................................................469

22 Toxicology Todd M. O’Hara and Thomas J. O’Shea

Introduction ............................................................................................471 Classes of Toxicants...............................................................................477 Elements .................................................................................................478 Mercury .........................................................................................478 Cadmium ......................................................................................480 Lead ...............................................................................................481 Organotins.....................................................................................481 Other Elements.............................................................................482 Halogenated Organics ............................................................................482 Accumulation and Variability .....................................................482 Organochlorine Pesticides and Metabolites ...............................484 Polychlorinated Biphenyls ...........................................................485 Other Organohalogens .................................................................487 Effects of Organochlorines on Metabolism ................................488 Effects of Organochlorines on Reproduction and Endocrine Function ....................................................................................490 Effects of Organochlorines on Immunocompetence and Epizootics ...........................................................................491 Biotoxins .................................................................................................493 Brevetoxin .....................................................................................493 Paralytic Shellfish Poisoning .......................................................494 Domoic Acid.................................................................................495 Ciguatera .......................................................................................496 Oil............................................................................................................496


Treatment and Diagnostic Procedures..................................................499 Dose Scaling..................................................................................499 Treatment......................................................................................499 Diagnosis .......................................................................................501 Acknowledgments ..................................................................................502 References ...............................................................................................502

23 Noninfectious Diseases Frances M. D. Gulland, Linda J. Lowenstine, and Terry R. Spraker

Introduction ............................................................................................521 Congenital Defects.................................................................................521 Neoplasia ................................................................................................522 Trauma ....................................................................................................522 Intraspecific Trauma ....................................................................522 Interspecific Trauma ....................................................................528 Anthropogenic Trauma ................................................................530 Miscellaneous .........................................................................................531 Integumentary System .................................................................531 Musculoskeletal and Dental Systems.........................................532 Respiratory System.......................................................................533 Digestive System ..........................................................................533 Genitourinary System ..................................................................534 Endocrine System .........................................................................535 Cardiovascular System.................................................................535 Lymphoid System .........................................................................536 Nervous System and Special Senses ...........................................536 Acknowledgments ..................................................................................537 References ...............................................................................................537

Section V

Diagnostic Imaging in Marine Mammals

24 Overview of Diagnostic Imaging William Van Bonn and Fiona Brook

Introduction ............................................................................................551 Imaging Science......................................................................................551 From Human to Marine Mammal Diagnostic Imaging ......................552 Application of Diagnostic Imaging Techniques...................................554 Conclusion ..............................................................................................555 Acknowledgments ..................................................................................556


25 Radiology, Computed Tomography, and Magnetic Resonance Imaging

William Van Bonn, Eric D. Jensen, and Fiona Brook

Introduction ............................................................................................557 Indications ..............................................................................................561 Limitations .............................................................................................565 Technique................................................................................................568 Clinical Applications .............................................................................574 Dolphin .........................................................................................574 Normal Radiographic Anatomy.......................................574 Radiographic Pathology....................................................579 Pinniped ........................................................................................581 Normal Radiographic Anatomy.......................................581 Radiographic Pathology....................................................585 Computed Tomographic Anatomy .......................................................586 Magnetic Resonance Imaging Anatomy, Dolphin ...............................587 Acknowledgments ..................................................................................588 References ...............................................................................................590

26 Ultrasonography Fiona Brook, William Van Bonn, and Eric D. Jensen

Introduction ............................................................................................593 Indications ..............................................................................................593 Limitations .............................................................................................594 Technique................................................................................................594 Equipment and Preparation .........................................................594 Image Orientation ........................................................................595 Clinical Applications .............................................................................596 Thoracic Imaging..........................................................................596 Heart and Mediastinum ...............................................................596 Lungs .............................................................................................597 Thoracic Lymph Nodes................................................................600 Abdominal Imaging ......................................................................601 Liver and Biliary System..............................................................601 Spleen ............................................................................................604 Pancreas.........................................................................................605 Gastrointestinal Tract ..................................................................605 Urinary Tract ................................................................................609 Reproductive Tract .......................................................................611 Males .................................................................................611 Females .............................................................................612


Eyes................................................................................................616 Musculoskeletal System ..............................................................616 Body Condition.............................................................................618 Conclusion ..............................................................................................618 Acknowledgments ..................................................................................618 References ...............................................................................................618

27 Flexible and Rigid Endoscopy in Marine Mammals Samuel R. Dover and William Van Bonn

Introduction ............................................................................................621 Indications ..............................................................................................622 Limitations .............................................................................................623 Equipment...............................................................................................624 Flexible Endoscopes......................................................................624 Rigid Telescopes ...........................................................................626 Light Sources ................................................................................626 Accessories and Instruments .......................................................627 Cameras.........................................................................................629 Video Monitors and Recorders ....................................................630 Clinical Applications in Cetaceans ......................................................630 Cetacean Gastroscopy ..................................................................630 Colonoscopy..................................................................................633 Respiratory Endoscopy .................................................................633 Urogenital .....................................................................................635 Clinical Applications in Other Marine Mammals ..............................635 Minimally Invasive Surgical Techniques .............................................636 Insufflation ....................................................................................636 Access............................................................................................637 Trocars and Cannulas...................................................................638 Closure ..........................................................................................639 Minimally Invasive Surgery in Cetaceans..................................640 Minimally Invasive Surgery in Other Marine Mammals..........640 Acknowledgments ..................................................................................641 References ...............................................................................................641

28 Thermal Imaging of Marine Mammals Michael T. Walsh and Edward V. Gaynor

Introduction ............................................................................................643 Technique................................................................................................643 History ....................................................................................................644 Cameras ..................................................................................................645


Clinical Applications .............................................................................645 Manatees .......................................................................................646 Pinnipeds .......................................................................................646 Cetaceans ......................................................................................647 Other Marine Mammal Species ..................................................649 Web Sites.......................................................................................650 Conclusion ..............................................................................................651 References ...............................................................................................651

Section VI

Medical Management of Marine Mammals

29 Marine Mammal Anesthesia Martin Haulena and Robert Bruce Heath

Introduction ............................................................................................655 Anesthetic Protocol................................................................................655 Preanesthetic Examination ..........................................................655 Choice of a Specific Anesthetic Protocol ...................................656 Monitoring Techniques..........................................................................656 Noninvasive Techniques..............................................................657 Invasive Techniques .....................................................................657 Support ....................................................................................................657 Cetaceans ................................................................................................657 Induction .......................................................................................657 Intubation......................................................................................660 Inhalation Anesthesia ..................................................................660 Monitoring ....................................................................................660 Support ..........................................................................................661 Emergencies ..................................................................................662 Otariids ...................................................................................................662 Induction .......................................................................................662 Intubation......................................................................................666 Inhalation Anesthesia ..................................................................667 Monitoring ....................................................................................668 Support ..........................................................................................668 Emergencies ..................................................................................669 Phocids ....................................................................................................670 Induction .......................................................................................670 Intubation......................................................................................674 Inhalation Anesthesia ..................................................................675 Monitoring ....................................................................................675 Support ..........................................................................................675 Emergencies ..................................................................................676


Odobenids ...............................................................................................677 Induction .......................................................................................677 Intubation and Inhalation Anesthesia ........................................680 Monitoring ....................................................................................680 Support ..........................................................................................680 Emergencies ..................................................................................681 Sirenians..................................................................................................681 Sea Otters................................................................................................681 Induction .......................................................................................681 Intubation......................................................................................683 Inhalation Anesthesia ..................................................................683 Monitoring ....................................................................................683 Support ..........................................................................................683 Emergencies ..................................................................................684 Ursids ......................................................................................................684 Conclusion ..............................................................................................684 Acknowledgments ..................................................................................684 References ...............................................................................................684

30 Intensive Care Michael T. Walsh and Scott Gearhart

Introduction ............................................................................................689 Records and Instructions .......................................................................689 Patient Evaluation..................................................................................689 Rehydration ............................................................................................690 Blood Transfusion...................................................................................692 Nutritional Therapy...............................................................................693 Hypoglycemia ...............................................................................693 Emaciation ....................................................................................693 Appetite Stimulants .....................................................................694 Respiratory Emergencies........................................................................695 Trauma ....................................................................................................695 Wound Management ..............................................................................696 Central Nervous System........................................................................696 Reproductive Emergencies.....................................................................697 Dystocia ........................................................................................697 Other Reproductive Emergencies................................................698 Antibiotics ..............................................................................................698 Analgesics ...............................................................................................699 Miscellaneous Therapeutic Agents.......................................................699 Support Equipment ................................................................................699 Conclusion ..............................................................................................700 References ...............................................................................................700


31 Pharmaceuticals and Formularies Michael K. Stoskopf, Scott Willens, and James F. McBain

Introduction ............................................................................................703 Routes for Administering Drugs to Marine Mammals .......................704 Dose Scaling ...........................................................................................705 Drug Interactions ...................................................................................705 Cimetidine and Antacids .............................................................705 Tetracyclines .................................................................................706 Fluoroquinolones ..........................................................................706 Other Antibiotics .........................................................................707 Antifungals....................................................................................708 Antiparasitic Drugs ......................................................................708 Steroids..........................................................................................708 Diuretics........................................................................................708 Drug Dosages..........................................................................................709 Acknowledgments ..................................................................................722 References ...............................................................................................722

32 Euthanasia Leah L. Greer, Janet Whaley, and Teri K. Rowles

Introduction ............................................................................................729 Stranded Animals ...................................................................................729 Display and Collection Animals...........................................................730 Methods of Euthanasia ..........................................................................730 Injectable Agents ....................................................................................731 Route of Administration..............................................................731 Barbiturates ...................................................................................732 Etorphine.......................................................................................732 T-61................................................................................................733 Paralytics .......................................................................................733 Inhalants .................................................................................................734 Physical Methods ...................................................................................734 Ballistics ........................................................................................734 Explosives......................................................................................736 Verification of Death..............................................................................736 Carcass Disposal.....................................................................................736 Acknowledgments ..................................................................................737 References ...............................................................................................737


Section VII

Marine Mammal Well-Being

33 U.S. Federal Legislation Governing Marine Mammals Nina M. Young and Sara L. Shapiro

Federal Legislation and Regulations—Discussion ...............................741 Introduction ..................................................................................741 The Responsible Regulating Agencies ........................................742 The Endangered Species Act........................................................743 Listing, Critical Habitat, and Recovery Plans................744 Protection for Listed Species ...........................................744 Permits ..............................................................................745 Consultations ....................................................................745 Enforcement ......................................................................745 Implementation of the Convention on International Trade in Endangered Species of Wild Fauna and Flora ........................................................................746 The Marine Mammal Protection Act .........................................750 The MMPA Moratorium on Taking................................750 Exemptions and Permits for Incidental Take.................750 Reauthorizations of the MMPA.......................................753 Marine Mammal Strandings and Health ........................753 The Animal Welfare Act..............................................................755 The Law.............................................................................755 Licensing and Registration...............................................755 Research Facilities ............................................................755 AWA Enforcement ............................................................755 Regulations........................................................................756 Space Requirements .........................................................756 Overlap among the Agencies and the Various Laws .....757 The Lacey Act of 1901.................................................................758 The Fur Seal Act...........................................................................758 Conclusion ....................................................................................758 Definitions and Abbreviations Pertaining to U.S. Marine Mammal Legislation ................................................759 Contact Information.........................................................762 Marine Mammal Permits: Frequently Asked Questions (FAQs)........762 The Marine Mammal Stranding Networks ................................762 Scientific Research and Enhancement Permits .........................763 Public Display Permits ................................................................764 Other Permits ...............................................................................765


Acknowledgments ..................................................................................765 References ...............................................................................................766

34 Public Health Daniel F. Cowan, Carol House, and James A. House

Introduction ............................................................................................767 Viral Infections .......................................................................................768 Poxviruses .....................................................................................768 Calicivirus.....................................................................................768 Influenza........................................................................................769 Rabies ............................................................................................769 Bacterial Infections.................................................................................769 Vibrio spp. .....................................................................................769 Edwardsiella spp. .........................................................................770 Clostridium spp............................................................................770 Leptospira......................................................................................770 Streptococcus ................................................................................770 Brucella .........................................................................................771 Erysipelothrix rhusiopathiae .......................................................771 Mycobacterium spp......................................................................771 Coxiella burnetii ..........................................................................772 Other Mixed Infections................................................................772 Mycoplasma Infections ..........................................................................772 Fungal Infections ....................................................................................773 Protozoal Infections ...............................................................................773 Toxoplasma gondii .......................................................................773 Cryptosporidium spp. ..................................................................774 Giardia spp. ..................................................................................774 Potential for Transmission of Infectious Disease from Marine Mammals to Humans..................................................774 Acknowledgments ..................................................................................775 References ...............................................................................................775

35 Water Quality Kristen D. Arkush

Introduction ............................................................................................779 Environmental Considerations..............................................................779 Space..............................................................................................780 System Water Source ...................................................................780 Temperature ..................................................................................780


Lighting .........................................................................................781 Salinity and pH.............................................................................781 Filtration .................................................................................................781 Microorganisms (as Pathogens and/or Indicators of Water Quality) ......................................................................783 Mechanisms of Sterilization..................................................................784 Ozone ............................................................................................785 Conclusions ............................................................................................786 Acknowledgments ..................................................................................786 References ...............................................................................................787

36 Nutrition and Energetics Graham A. J. Worthy

Introduction ............................................................................................791 Energy Requirements .............................................................................791 Metabolic Rate..............................................................................792 Thermoregulation.........................................................................794 Locomotion ...................................................................................796 Summary: Average Daily Metabolic Rate ..................................799 Water Requirements.....................................................................799 Fasting and Starvation..................................................................801 The Bioenergetic Scheme ......................................................................803 Maintenance Energy.....................................................................804 Production Energy ........................................................................804 Reproduction .....................................................................804 Molt ...................................................................................807 Heat Increment of Feeding ..........................................................807 Fecal and Urinary Energy Losses ................................................809 Calculation of Gross Energy Requirements ...............................810 Prey..........................................................................................................811 Species That Marine Mammals Consume in Captivity and in the Wild.........................................................................811 Seasonal Changes in Prey Composition .....................................813 Major Nutritional Disorders..................................................................813 Thiamine Deficiency....................................................................813 Hyponatremia ...............................................................................814 Vitamins A, D, and E ...................................................................815 Vitamin C......................................................................................816 Scombroid Poisoning....................................................................816 Conclusions ............................................................................................817 Acknowledgments ..................................................................................817 References ...............................................................................................817


37 Hand-Rearing and Artificial Milk Formulas Forrest I. Townsend, Jr. and Laurie J. Gage

Introduction ............................................................................................829 Cetaceans ................................................................................................829 Formula .........................................................................................829 Delivery Methods and Techniques .............................................830 Feeding Frequency and Daily Requirements..............................830 Monitoring Neonates ...................................................................831 Weaning Procedures .....................................................................831 Other Practical Information ........................................................831 References and Suggested Further Reading ................................831 Pinnipeds.................................................................................................832 Harbor Seals ..................................................................................832 Formula .............................................................................832 Delivery Methods and Techniques..................................833 Feeding Frequency and Daily Requirements ..................833 Weaning Procedures..........................................................834 Other Practical Information.............................................834 References and Suggested Further Reading ....................834 Elephant Seals...............................................................................836 Formulas............................................................................836 Fish Mash ..........................................................836 Elephant Seal Formula......................................836 ESF 50–50 ..........................................................836 ESF 75–25...........................................................837 Feeding Frequency and Daily Requirements ..................837 Delivery Methods and Techniques..................................838 Weaning Procedures..........................................................838 Other Practical Information.............................................838 References and Suggested Further Reading ....................838 Sea Lions .................................................................................................839 Formula .........................................................................................839 Delivery Methods and Techniques .............................................839 Feeding Frequency and Daily Requirements..............................840 Weaning Procedures .....................................................................840 Other Practical Information ........................................................840 References and Suggested Further Reading ................................840 Walruses ..................................................................................................841 Formulas........................................................................................841 Beginning Formula............................................................841 Maintenance formula .......................................................841 Feeding Frequency and Daily Requirements..............................842


Delivery Methods and Techniques .............................................842 Weaning Procedures .....................................................................842 Other Practical Information ........................................................842 References and Suggested Further Reading ................................842 Manatees .................................................................................................843 Formulas........................................................................................843 Miami Seaquarium Formula ............................................843 SeaWorld Formula ............................................................843 Delivery Methods and Techniques .............................................843 Feeding Frequency and Daily Requirements..............................844 Weaning Procedures .....................................................................844 Other Practical Information ........................................................844 References and Suggested Further Reading ................................845 Sea Otters................................................................................................845 Formula and Preparation .............................................................845 Delivery Methods and Techniques .............................................845 Feeding Frequency and Daily Requirements..............................846 Weaning Procedures .....................................................................846 Other Practical Information ........................................................846 References and Suggested Further Reading ................................847 Polar Bears ..............................................................................................847 Formulas........................................................................................847 Delivery Methods and Techniques .............................................848 Feeding Frequency and Daily Requirements..............................848 Weaning Process ...........................................................................848 Other Practical Information ........................................................848 References and Suggested Further Reading ................................848 Acknowledgments ..................................................................................849

38 Tagging and Tracking Michelle E. Lander, Andrew J. Westgate, Robert K. Bonde, and Michael J. Murray

Introduction ............................................................................................851 Tracking Methodologies: A Brief Overview.........................................851 Pinnipeds.................................................................................................857 Cetaceans ................................................................................................862 Manatees .................................................................................................866 Sea Otters................................................................................................870 Polar Bears ..............................................................................................874 Conclusion ..............................................................................................874 Acknowledgments ..................................................................................874 References ...............................................................................................874


39 Marine Mammal Transport Jim Antrim and James F. McBain

Introduction ............................................................................................881 Regulations .............................................................................................881 History of Marine Mammal Transport.................................................882 Cetaceans ......................................................................................882 Pinnipeds .......................................................................................888 Sea Otters......................................................................................888 Sirenians ........................................................................................889 Polar Bears.....................................................................................889 Additional Medical Considerations ......................................................889 Conclusion ..............................................................................................890 Acknowledgments ..................................................................................891 References ...............................................................................................891

Section VIII

Specific Medicine and Husbandry of Marine Mammals

40 Cetacean Medicine James F. McBain

Introduction ............................................................................................895 Philosophy ..............................................................................................895 Clinical Examination .............................................................................896 History...........................................................................................896 Visual Examination ......................................................................897 How Does the Animal Feel? .......................................................897 Buoyancy .......................................................................................897 Decreased Buoyancy .........................................................898 Increased Buoyancy ..........................................................898 Listing ................................................................................898 Social Behavior .............................................................................898 Hands-On Examination................................................................899 Urine Collection...........................................................................899 Stool Samples................................................................................899 Milk Samples ................................................................................899 Blowhole........................................................................................900 Additional Diagnostic Aids ...................................................................900 Body Weight ..................................................................................900 Ultrasonography ...........................................................................900 Radiography ..................................................................................900 Clinical Laboratory Tests.............................................................900


Clinical Pathology..................................................................................901 Case Example: Pulmonary Disease .............................................901 Indicators of Inflammatory Disease ................................901 Therapeutics ...........................................................................................903 Surgery...........................................................................................903 Medical Therapy...........................................................................903 Oral Route.........................................................................903 Subcutaneous Route .........................................................904 Intramuscular Route.........................................................904 Intravenous Route ............................................................904 Topical Route ....................................................................904 Final Thoughts .......................................................................................905 Acknowledgments ..................................................................................905 References ...............................................................................................905

41 Seals and Sea Lions Frances M. D. Gulland, Martin Haulena, and Leslie A. Dierauf

Introduction ............................................................................................907 Husbandry...............................................................................................907 Pools, Haul-Out Areas, and Enclosures ......................................907 Feeding ..........................................................................................908 Restraint..................................................................................................908 Physical Restraint.........................................................................908 Mechanical Restraint ...................................................................909 Chemical Restraint ......................................................................909 Physical Examination ............................................................................909 Diagnostic Techniques...........................................................................910 Blood Collection ...........................................................................910 Urine..............................................................................................910 Cerebrospinal Fluid ......................................................................911 Biopsies..........................................................................................911 Therapeutic Techniques ........................................................................911 Topical ...........................................................................................911 Oral................................................................................................911 Aerosol ..........................................................................................912 Subcutaneous ................................................................................912 Intramuscular................................................................................912 Intravenous ...................................................................................912 Intraosseous ..................................................................................912 Intraperitoneal ..............................................................................912 Diseases...................................................................................................913 Integumentary System .................................................................913 Musculoskeletal System ..............................................................915


Digestive System ..........................................................................916 Respiratory System.......................................................................917 Cardiovascular ..............................................................................919 Urogenital System ........................................................................919 Endocrine System .........................................................................920 Eyes................................................................................................920 Nervous System............................................................................921 Acknowledgments ..................................................................................922 References ...............................................................................................922

42 Walruses Michael T. Walsh, Brad F. Andrews, and Jim Antrim

Introduction ............................................................................................927 Biology.....................................................................................................927 Reproduction ..........................................................................................928 Diet..........................................................................................................929 Physical Examination ............................................................................929 Restraint..................................................................................................930 Manual ..........................................................................................930 Sedation and General Anesthesia................................................930 Specimen Collection and Diagnostic Techniques ...............................930 Medical Problems...................................................................................931 Dermatology .................................................................................931 Ophthalmology .............................................................................932 Tusk Infections and Trauma........................................................933 Foreign Bodies...............................................................................934 Intestinal Disease .........................................................................934 Miscellaneous Diseases................................................................935 Acknowledgments ..................................................................................935 References ...............................................................................................935

43 Manatees Gregory D. Bossart

Introduction ............................................................................................939 Natural History ......................................................................................939 Anatomy, Physiology, and Behavior .....................................................941 Husbandry...............................................................................................942 Habitat Requirements ..................................................................942 Water Requirements.....................................................................942 Nutrition .......................................................................................943 Restraint, Handling, and Transport ............................................944


Physical Examination...................................................................946 Diagnostic Techniques .................................................................946 Therapeutics .................................................................................948 Anesthesia .....................................................................................950 Environmental Diseases ........................................................................951 Brevetoxicosis ...............................................................................951 Cold Stress Syndrome ..................................................................951 Infectious Diseases.................................................................................952 Parasites ........................................................................................952 Miscellaneous Conditions .....................................................................953 Neoplasia.......................................................................................953 Neonatal Disease..........................................................................953 Human-Related Traumatic Injuries ............................................954 Acknowledgments ..................................................................................958 References ...............................................................................................958

44 Sea Otters Pamela Tuomi

Introduction ............................................................................................961 History ....................................................................................................961 Classification ..........................................................................................962 Anatomy .................................................................................................963 Vision ......................................................................................................965 Social Organization ................................................................................965 Reproduction ..........................................................................................965 Causes of Mortality in Free-Living Otters ...........................................967 Feeding and Metabolism........................................................................967 Husbandry...............................................................................................969 Captive Nutrition...................................................................................971 Physical and Chemical Restraint..........................................................971 Clinical Examination .............................................................................973 Medical Abnormalities ..........................................................................974 Hypoglycemia ...............................................................................974 Hyperthermia................................................................................974 Hypothermia .................................................................................975 Loss of Coat Condition ................................................................975 Oil Exposure .................................................................................976 Abnormalities of Clinical Chemistry .........................................977 Gastroenteritis ..............................................................................978 Parasites ........................................................................................978 Miscellaneous Conditions ...........................................................979 Surgery ....................................................................................................979 Dentistry .................................................................................................980


Preventive Medicine ..............................................................................980 Acknowledgments ..................................................................................980 References ...............................................................................................980

45 Polar Bears Michael Brent Briggs

Introduction ............................................................................................989 Natural History and Physiology ...........................................................989 Nutrition .................................................................................................990 Nutrition of Juveniles, Early Pregnant, and Lactating Females..............................................................991 Infants............................................................................................992 Geriatrics.......................................................................................992 Reproduction ..........................................................................................992 Endocrinology .........................................................................................992 Reproductive Hormones ..............................................................992 Thyroid Hormones .......................................................................993 Housing ...................................................................................................993 Behavior ..................................................................................................994 Physical Examination ............................................................................994 Venipuncture ..........................................................................................995 Mechanical or Manual Restraint ..........................................................996 Anesthesia...............................................................................................996 Ketamine .......................................................................................997 Ketamine/Xylazine ......................................................................998 Tiletamine HCl and Zolazepam HCl .........................................998 Telazol/Medetomidine .................................................................999 Etorphine.......................................................................................999 Carfentanil ....................................................................................999 Fentanyl Citrate............................................................................999 Inhalation Agents .........................................................................999 Systemic Diseases ................................................................................1000 Developmental/Anomalous Diseases .......................................1000 Nutritional Diseases ..................................................................1000 Neoplasia.....................................................................................1000 Infectious Diseases .....................................................................1001 Viral Disease ...................................................................1001 Bacterial Disease.............................................................1001 Mycotic Disease..............................................................1001 Parasitic Disease .............................................................1002 Skin Disease................................................................................1002 Dental Disease............................................................................1003 Trauma ........................................................................................1003 Toxins ..........................................................................................1003


Zoonoses ...............................................................................................1003 Acknowledgments ................................................................................1003 References .............................................................................................1004

Appendices Appendix A Conversions ...............................................................1011 Appendix B Abbreviations ............................................................1015 Appendix C Characteristics of Common Disinfectants ....1017 Index ...........................................................................................................1019


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I Emerging Pathways in Marine Mammal Medicine



1 Marine Mammals as Sentinels of Ocean Health Michelle Lynn Reddy, Leslie A. Dierauf, and Frances M. D. Gulland

Introduction It was January 1958, when Rachel Carson, a marine biologist who had been working with the U.S. Fish and Wildlife Service, received a letter from Olga Owens Huckins of Duxbury, Massachusetts. The letter told of birds dying after local applications of the pesticide DDT (dichlorodiphenyl trichloroethane) (Gore, 1994). DDT had already been known to have detrimental effects on birds (Robbins et al., 1951), and the evidence would continue to grow (Robinson, 1969; Faber and Hickey, 1973; Fry and Toone, 1981). More sensitive to the pesticides in their environment, the birds showed effects long before effects were seen in other wildlife species or in humans. Rachel Carson went on to write the landmark book Silent Spring (Carson, 1962), alerting the general public to the insidious effects of chemical pollutants. People were becoming better at understanding the importance of recognizing adverse reactions of wildlife to anthropogenic hazards in the environment. Carson’s local birds were sentinels of environmental changes that in time were shown to affect human health. However, these were not the first avian sentinels. At the turn of the 20th century, experiments by the Bureau of Mines showed that canaries taken into mines collapsed when exposed to carbon monoxide gas (the birds recovered when exposed to fresh air). Miners were able to avoid possible disaster by carrying caged canaries with them into mineshafts and tunnels. The birds alerted them to the presence of the deadly invisible gas (Burrell and Seibert, 1916).

Sentinels The word sentinel has its origins in the Latin, sentire, which means to perceive or feel (Morris, 1975), and is now used to mean a person or animal who guards the group against surprise. The National Research Council (1991) defines an animal sentinel system as “a system in which data on animals exposed to contaminants in the environment are regularly and systematically collected and analyzed to identify potential health hazards to other animals or humans.” Sentinel systems provide knowledge needed to facilitate early responses to potentially hazardous conditions and to allow for more effective resource management. For such systems to be effective in controlling and preventing disease, they must be simple, sensitive, representative, 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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CRC Handbook of Marine Mammal Medicine

and timely (CDC, 1988). Ideally, sentinels should detect changes prior to their effects becoming irreversible. Depending on what these systems are designed to monitor, animal sentinels can be wild or domestic, maintained in a laboratory or at a zoological park, and they can be terrestrial or marine (National Research Council, 1991). Animal species that are “charismatic megafauna”—such as whales and dolphins—make particularly good sentinels, because they have special public appeal and can be more effective at drawing societal attention and action to the plight of ecosystems. Invertebrates such as bivalves (clams, mussels, oysters) have been used widely as bioindicators of environmental contamination (Butler, 1973; Farrington et al., 1983). Bivalves are sedentary with relatively stable populations, so body burdens of contaminants reflect local conditions and can be used for long- and short-term pollution assessment. Additionally, they have a universal distribution that facilitates data comparison between many regions; they concentrate contaminants in their tissues; they have little or no detectable reactive enzyme systems to metabolize toxins, which makes assessment reasonably accurate; they are relatively tolerant of polluted conditions; and they are commercially available worldwide and thus have public health implications (Farrington et al., 1983; National Research Council, 1991). Vertebrates are also used as sentinels, and because they are at higher trophic levels than invertebrates, they are more likely to show the biomagnification effects of contaminants. Contaminant effects on sentinels, whether invertebrate or vertebrate, may occur at the suborganismal, organismal, or population level (Keith, 1996). Suborganismal effects include genotoxic effects, alterations in enzyme function, metallothionein induction, changes in thyroid function and retinol homeostasis, and hematological changes. Effects at the organismal level include pathological lesions, and alterations in development, growth, reproduction, and survival. Effects at the population level include alterations in abundance and distribution and changes in species assemblages (McCarthy and Shugart, 1990).

Ecosystem Changes Detected by Sentinels Canaries are no longer used in mines; modern, technological carbon monoxide detection and monitoring devices have replaced them. Today the scope of environmental concern has expanded. The great number of humans inhabiting the Earth, in concert with their ever-increasing consumption and destruction of resources, places enormous pressures on the environment. By 2010, it is predicted that the Earth’s population will be 9.3 billion (Colborn et al., 1996). Yet we are far from understanding the effects of the alterations we are imposing on our environment. However, if data are carefully collected and analyzed from properly designed, implemented, and coordinated animal sentinel programs, we can make important inroads in detecting and mitigating some of the environmental threats we are inadvertently imposing upon ourselves. The effects of humans can be found in every ecosystem, whether it is deep in the dampest rain forest, high on the most frigid mountain top, or surrounded by the driest desert. However, the habitat that defines the planet Earth is the ocean, which covers 79% of the Earth’s surface. These effects may be direct, such as by the overharvesting of commercial fisheries, or indirect, through effects of runoff and global warming. Oceans facilitate the distribution of potentially toxic contaminants such as heavy metals and organochlorine (OC) chemicals. Comprising industrial chemicals such as polychlorinated biphenyls (PCBs) and chlorinated pesticides such as DDT, OCs tend to be stable and lipophilic. A group of experts attending a meeting on “Chemically Induced Alterations in Sexual Development: The Wildlife/Human Connection” concurred that “we are certain of the following: A large number of man-made chemicals that have been released into the environment … have the potential to disrupt the endocrine system of animals, including humans” (Colborn and Clement, 1992).


Marine Mammals as Sentinels of Ocean Health

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In the sea, contaminants in runoff from urban, industrial, and agricultural activities intermix and bioaccumulate up the food chain, attaining the greatest concentrations in animals at the highest trophic levels, such as marine mammals. At an international workshop on marine mammals and persistent ocean contaminants in 1998, invited experts concluded that “there is good reason to be concerned that survival and reproduction in certain marine mammal populations may have been affected, and are being affected, by persistent contaminants, particularly OCs.” The workshop also concluded that there is a need for multidisciplinary studies on the significance of ocean contaminants in relation to the health and well-being of marine mammals (Marine Mammal Commission, 1998; see Chapter 22, Toxicology). Activities of humans and terrestrial animals also impact ocean health in other ways. Recently identified pathogens in marine mammals, such as Giardia lamblia, Sarcocystis neurona, Toxoplasma gondii, and antibiotic-resistant enteric bacteria, may all originate in waste from humans or their activities (Buergelt and Bonde, 1983; Olsen et al., 1997; Parveen et al., 1997; Johnson et al., 1998; LaPointe et al., 1999; Measures and Olsen, 1999). Runoff also increases nutrient load and availability, enhancing blooms of potentially toxic marine algae species such as Alexandrium spp. (produce saxitoxins), Gymnodinium breve (Ptychodiscus brevis) (produce brevitoxin), and Pseudonitzschia australis (produce domoic acid) (Geraci and Lounsbury, 1993; Smolowitz and Doucette, 1995; Scholin et al., 2000; see Chapter 2, Emerging and Resurging Diseases; Chapter 22, Toxicology). Whether such infectious agents and algal blooms are increasing in prevalence or are merely being detected more readily due to increasing awareness of ocean and marine mammal health issues is still subject of debate (Harvell et al., 1999). The ocean is also a sink for excess heat, and as such, it is an effective global thermostat (Carson, 1951). The National Oceanic and Atmospheric Administration (NOAA) National Climatic Data Center (NCDC) tracks land and sea temperature measurements. On its Web site (http://www.ncdc.noaa.gov/ol/climate/globalwarming.html), the NCDC reports that global surface temperatures have increased about 1°F (0.3 to 0.6°C) since the late 19th century, and about 0.5°F (0.2 to 0.3°C) over the past 40 years, which is the period with the most credible data. This warming trend is due to what is commonly known as the greenhouse gas effect—a result of industrial output of carbon dioxide, methane, and nitrous oxide that accumulates in the atmosphere and traps heat. Global climate change may alter animal abundance, distribution, and migration patterns, and has the potential to influence disease patterns worldwide (Aguilar and Raga, 1993; Daszak et al., 2000). Potential effects on cetaceans are reviewed by Burns (2000). Another form of pollution is noise pollution. Cetaceans have drawn attention to the increase in noise levels in the oceans (Richardson et al., 1995; National Research Council, 2000). Cetaceans use sound for a variety of purposes including foraging, communication, and navigation. It is feared that low-frequency, high-intensity noise generated by maritime shipping, polar icebreakers, offshore drilling, seismic surveys, oceanographic testing, and military use in the world’s oceans is a potentially serious problem for cetaceans, so there is a critical need for data on cetacean hearing for assessing the effects of such noise on these animals. Sound sources that have been developed for use in monitoring changes in ocean temperatures and detecting stealth submarines are currently hot topics. These sounds travel long distances, perhaps even masking sounds produced by marine mammals (National Research Council, 2000).

Marine Mammals as Sentinels Holden (1972) was perhaps the first to formally propose the use of marine mammals as environmental sentinels. Marine mammals are good indicators of mid- to long-term changes in the environment, because many species have long life spans, feed at or near the top of the food chain, and have extensive fat stores (Aguilar and Borrell, 1994). Ironically, the blubber


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that plays a crucial role in nutrition, buoyancy, and thermoregulation for these animals is an ideal repository for some contaminants. While the most inert and lipophilic of these contaminants may remain stored in the blubber until the animal dies, others may be metabolized, especially in times of physiological challenge such as illness, extreme temperature, nutritional compromise, or pregnancy and lactation (DeFreitas et al., 1969; McKenzie et al., 1997). The California sea lion (Zalophus californianus), harbor seal (Phoca vitulina), bottlenose dolphin (Tursiops truncatus), and beluga (Delphinapterus leucas) have been identified as model species for investigations into the effects of environmental contaminants on marine mammals (Marine Mammal Commission, 1998). The ecology and life histories of these animals are relatively well studied, they are relatively common thus more readily sampled, and they are well represented in facilities where breeding programs have been successful (Andrews et al., 1997). One way to more accurately ascertain contaminant effects on wild marine mammal populations is to use biomarkers in samples carefully collected from free-ranging animals (Peakall, 1992; Aguilar and Borrell, 1994). This is particularly true if samples are collected from representative members of populations that are the focus of long-term monitoring programs (Gaskin et al., 1982; Scott et al., 1990; Addison and Smith, 1998; Addison et al., 1998), especially when relevant biological data and health histories are available (Scott et al., 1990). However, regulations often prohibit collecting samples from young and their accompanying mothers in the wild, and there is no guarantee that any particular individual will be available for sampling. Additionally, data can be affected by variation in sample collection, handling, and processing, which can be difficult to control under field conditions. For example, when collecting blubber biopsies, it may be difficult to regulate the location and depth of the biopsy, both of which may affect results depending on the species (Aguilar and Borrell, 1994). In addition, because of the logistical difficulties and expenses involved in such operations, few are undertaken. Hunted marine mammals, such as the bowhead whale (Balaena mysticetus) harvested by the Inuit in Alaska, can also be sampled to yield information on ocean contaminants and marine mammal health (O’Hara et al., 1999). Because these animals are freshly dead and can be examined in detail, levels of contaminants can be correlated with histological changes in individual animals. Because bowhead whale populations have been well monitored, contaminant data from individuals yield insight into changes in reproduction and survival at the population level. Marine mammals have helped draw public attention to the current plight of fish stocks. For example, the western population of Steller sea lions (Eumetopias jubatus) has declined by more than 70% since the 1970s (Ferrero and Fritz, 2000), resulting in the addition of this species to the federal list of endangered species (National Marine Fisheries Service, 1992). The cause of the decline remains unclear and may be a combination of factors. Management actions have been implemented to reduce potential interactions between Steller sea lions and the Alaskan groundfish fishery (Ferrero and Fritz, 2000). However, it has been hypothesized that the large-scale harvesting of fish and whales that occurred from the 1950s through the early 1970s in the Bering Sea and Gulf of Alaska (National Research Council, 1996) may have altered the food web, allowing walleye pollock (Theragra chalcogramma) to become a dominant fish species (see Bowen, 1997). Pollock is an economically significant fish, as well as an important prey item for Steller sea lions (Lowry et al., 1989), so shortage in pollock stocks could significantly contribute to the decreasing numbers of these pinnipeds. Understanding the size composition of fishes eaten by a predator such as the Steller sea lion in relation to those of the commercial catch can lend much insight into marine mammal–fisheries interactions (Frost and Lowry, 1986). The Steller sea lion may thus prove to be an important sentinel for fish stocks in the Bering Sea. The exceptional hearing and sound production capabilities of cetaceans have long been recognized by scientists. Many species can hear sounds well outside the range of human hearing



7

(Ridgway, 1997). Much has been learned about hearing in small cetacean species that are housed at marine mammal facilities. However, little or nothing is known about hearing in other cetacean species, such as the large baleen whales and some of the larger toothed whales such as beaked whales and the great sperm whale (Physeter catadon). Recently, intense sound from naval vessels has been implicated in several stranding events at various locations across the globe (Frantzis, 1998). Studies are currently under way to investigate the effects of anthropogenic noise on cetaceans (e.g., Au et al., 1999; Erbe and Farmer, 2000; Finneran et al., 2000; Schlundt et al., 2000). These studies will aid understanding of the effects of intense noise, which will contribute to the development of mitigation strategies ultimately to help find a balance between the basic needs of marine mammals and the important role the ocean plays in commerce, exploration, national defense, and travel. Stranded marine mammals are another source of information about the ocean environment (Geraci and Lounsbury, 1993; Gulland, 1999). Not only can they be sampled to quantify contaminant levels in tissues, but they can also alert researchers to diseases that are present in the more inaccessible wild animals that would be difficult to detect in random samplings of such populations. For example, 20% of sexually mature California sea lions that stranded and died along the northern coast of California showed neoplasia when examined post-mortem (Gulland et al., 1996). In comparison, only one case of neoplasia has been observed in California sea lions at rookeries on San Miguel Island, California, where more than 100,000 sea lions live (Spraker, pers. comm.). Study of neoplasia pathogenesis is more readily performed on stranded sea lions than on those in rookeries, and thus stranded animals essentially serve as sentinels for their wild conspecifics. Similarly, stranded belugas in the St. Lawrence estuary serve as sentinels of the health of the estuary. These whales have an unusually high prevalence of tumors and diseases for cetaceans, suggesting that this population is immunocompromised (Martineau et al., 1988; 1999; De Guise et al., 1994). These findings, coupled with the charismatic appeal of the beluga, have helped raise concern over contaminant levels in the St. Lawrence River and estuary. A number of infectious agents in marine mammals were first identified in stranded animals, after which their presence in the free-ranging population was confirmed. These include phocine distemper virus (PDV), which caused the death of over 18,000 harbor seals in Europe in 1988 (Osterhaus and Vedder, 1988), phocine herpes virus (PhHV1) isolated from stranded harbor seals in 1985 (Osterhaus et al., 1985), and Brucella in a variety of species (Ross et al., 1994; Garner et al., 1997) (see Chapter 15, Viral Diseases; Chapter 16, Bacterial Diseases). Live stranded animals offer an opportunity to monitor clinical signs that may result from changes in ocean health. For example, thorough examination of stranded, sick California sea lions resulted in the detection of domoic acid, a recently identified marine biotoxin, produced by the diatom Pseudonitzschia australis. The sea lions had consumed toxin-laden anchovies, and the domoic acid concentrated in the tissues of the sea lions caused muscle tremors, seizures, and death (Scholin et al., 2000) (see Chapter 2, Emerging and Resurging Diseases). In this case, the findings warned against human consumption of the anchovies, and increased monitoring of other seafood in the area. Stranded animals do not constitute an ideal sentinel system, as they do not represent the entire population (Aguilar and Borrell, 1994). In addition, samples of stranded animals are rarely age and sex structured, and biological data such as individual life histories, feeding habits, reproductive success, or disease progression are not typically available. Furthermore, contaminant levels in tissues collected from animals found dead may be significantly affected by decomposition of the samples (Borrell and Aguilar, 1990) (see Chapter 22, Toxicology). Marine mammals maintained at research and display facilities can be effective sentinels. The authors of the Marine Mammal Protection Act (MMPA), passed by Congress in 1972 (see Chapter 33, Legislation), understood the value of marine mammals in collections for conducting


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research and raising environmental awareness. They specifically allowed for the collection of marine mammals, stating “(3) there is inadequate knowledge of the ecology and population dynamics of such marine mammals and of the factors which bear upon their ability to reproduce themselves successfully; (4) negotiations should be undertaken immediately to encourage the development of international arrangements for research on, and conservation of, all marine mammals” (MMPA sec. 2, p. 2). Reijnders (1988) stated, “Even more than before, marine mammals in captivity should be used to obtain a set of reference data to interpret values obtained from animals expected to be affected by contaminants.” There are many advantages to using animals under human care as sentinels. Longitudinal health data are available for long-term studies and may provide insight into transgenerational and long-term health trends. These animals are fed wild-caught fish that have naturally occurring levels and mixtures of contaminants. These contaminants can be identified and quantified to provide insight not only into the dietary exposure of the marine mammals, but also into ecosystem levels and distribution of OCs that may impact the seafood-consuming public. In addition, tissues and fluids, including storage (blubber) and circulating (blood) compartments, can be regularly and systematically collected using conditioned husbandry behaviors, whereby the animals cooperate in specimen collection. Biological data such as age, sex, nutritional state, and reproductive and health histories can be recorded and correlated with measured contaminant levels. Changes in blubber levels can be correlated with levels in blood. Studies can be designed to establish effective biomarkers for monitoring complex physiological functions, such as immune and neurological responses and effects on reproduction. Contaminant monitoring is currently ongoing in San Diego where a large collection of bottlenose dolphins is maintained by the U.S. Navy. The animals reside in netted enclosures in San Diego Bay, California, often work in the open ocean, and are fed a diet from known sources. Preliminary research has revealed that preprandially collected blood can be used to estimate blubber levels of contaminants using lipid-normalized levels of OCs found in blood (Reddy et al., 1998). Milk samples collected voluntarily (Kamolnick et al., 1994) from lactating females in this population showed that from day 94 to day 615 of lactation, lipid-normalized levels of PCB and DDE (dichlorodiphenyl dichloroethylene) decreased by 69 and 82%, respectively (Ridgway and Reddy, 1995). In addition, preliminary data showed that concentrations of several OC contaminants in maternal blubber correlated strongly with reproductive outcome in these animals (Reddy et al., 2000). This population may provide a useful benchmark for marine mammal OC studies. Marine mammals can also be temporarily collected for contaminant studies; two such studies have been conducted with groups of harbor seals (Reijnders, 1986; Brouwer et al., 1989; de Swart et al., 1994; 1996; Ross et al., 1995; 1996). In these studies, half of the animals were fed fish from a highly polluted source and the other half were fed fish from a lesspolluted source. Results showed that animals fed higher levels of contaminants had reduced levels of circulating thyroid hormone and vitamin A, suppressed immune responses, and reduced reproductive success. A comprehensive marine mammal sentinel system would best include data collected from many sources including stranded animals, wild populations, and animals in collections. To ensure data quality, and to facilitate comparison between studies, it is important to standardize sample collection and handling protocols and to maintain archived samples to study as new analytical methods and technologies are developed (Wise et al., 1993) (see Chapter 21, Necropsy; Chapter 22, Toxicology). Linking these studies with laboratory toxicity studies should provide valuable insight into natural exposure and potential risk assessment and management strategies (National Research Council, 1991; Ross, 2000).



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Conclusion Marine mammals are effective ambassadors for the ocean environment because of their great public appeal. O’Shea points out, “For the general public, marine mammals are one of the most conspicuous components of marine biological diversity. Any that come ashore dead or ill raise the levels of uneasiness about the health of our oceans” (Geraci and Lounsbury, 1993). Stenciled images of marine mammals on storm drains in coastal cities with reminders of “No dumping, we live downstream” support this sentiment. More than ever, it is imperative to use an interdisciplinary and interagency approach. Long-term monitoring of populations and toxicological and disease investigations are expensive, time-consuming, and complex. The collaborative expertise of specialists, including oceanographers, geographers, chemists, biologists, physicians, veterinarians, epidemiologists, and pathologists, is needed to understand the effects of ocean health on the health of marine mammals and potentially humans. Klamer et al. (1991) predicted, “If the increase in ocean PCB concentrations continues, it may ultimately result in the extinction of fish-eating marine mammals.” But there is still time. The ocean has not yet fallen silent in the fashion forewarned by Rachel Carson in Silent Spring (1962). The great mammals of the sea have much to tell us, if only we learn to listen.

Acknowledgments The authors thank Gwen Griffith, Scott Newman, Andy Draper, and Donna Staples for reviewing this chapter.

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Burrell, G.A., and Siebert, F.M., 1916, Gases found in coal mines, Miners’ Circular 14, Bureau of Mines, Department of the Interior, Washington, D.C. Butler, P.A., 1973, Organochlorine residues in estuarine mollusks, 1965–1972: National Pesticide Monitoring Program, Pest. Monit. J., 6: 238–362. Carson, R.L., 1951, The Sea around Us, Oxford University Press, New York, 230 pp. Carson, R., 1962, Silent Spring, Houghton Mifflin Company, Boston, MA, 368 pp. CDC (Centers for Disease Control), 1988, Guidelines for Evaluating Surveillance Systems, MMWR, 37(S-5). Colborn, T., and Clement, C. (Eds.), 1992, Chemically Induced Alterations in Sexual and Functional Development: The Wildlife/Human Connection, Princeton Scientific Publishing, Princeton, NJ, 403 pp. Colborn, T., Dumanoski, D., and Myers, J.P., 1996, Here, there and everywhere, in Our Stolen Future: Are We Threatening Our Fertility, Intelligence and Survival? Dutton, New York, 132–133. Daszak, P., Cunningham, A.A., and Hyatt, A.D., 2000, Emerging infectious diseases of wildlife—threats to biodiversity and human health, Science, 287: 443–449. DeFreitas, A.S.W., Hart, J.S., and Morley, H.V., 1969, Chronic cold exposure and DDT toxicity, in Chemical Fallout: Current Research on Persistent Pesticides, Miller, M.W., and Berg, G.G. (Eds.), Charles C Thomas, Springfield, IL, 361–367. De Guise, S., Lagace, A., and Béland, P., 1994, Tumors in St. Lawrence beluga whales (Delphinapterus leucas), Vet. Pathol., 31: 444–449. de Swart, R.L., Ross, P.S., Vedder, L.J., Timmerman, H.H., Heisterkamp, S., Van Loveren, H., Vos, J.G., Reijnders, P.J.H., and Osterhaus, A.D.M.E., 1994, Impairment of immune function in harbor seals (Phoca vitulina) feeding on fish from polluted waters, Ambio, 23: 155–159. de Swart, R.L., Ross, P.S., Vos, J.G., and Osterhaus, A.D.M.E., 1996, Impaired immunity in harbor seals (Phoca vitulina) exposed to bioaccumulated environmental contaminants: Review of a long-term study, Environ. Health Perspect., 104 (Suppl. 4): 823–828. Erbe, C., and Farmer, D.M., 2000, A software model to estimate zones of impact on marine mammals around anthropogenic noise, J. Acoust. Soc. Am., 108: 1327–1331. Faber, R., and Hickey, J., 1973, Eggshell thinning, chlorinated hydrocarbons, and mercury in inland aquatic bird eggs, 1969 and 1970, Pest. Monit. J., 7: 27–36. Farrington, J.A., Goldberg, E.D., Risebrough, R.W., Martin, J.H., and Bowen, V.T., 1983, U.S. “Mussel Watch” 1976–1978: An overview of the tracemetal, DDE, PCB, hydrocarbon, and artificial radionuclide data, Environ. Sci. Technol., 17: 490–496. Ferrero, R., and Fritz, L., 2000, Steller sea lion/Alaskan groundfish fisheries interactions draw increased management attention, Mar. Mammal Soc. Newsl., 8: 2. Finneran, J.J., Schlundt, C.E., Carder, D.A., Clark, J.A., Young, J.A., Gaspin, J.B., and Ridgway, S.H., 2000, Auditory and behavioral responses of bottlenose dolphins (Tursiops truncatus) and a beluga whale (Delphinapterus leucas) to impulsive sounds resembling distant signatures of underwater explosions, J. Acoust. Soc. Am., 108: 417–431. Frantzis, A., 1998, Does acoustic testing strand whales? Nature, 392: 29. Frost, K.J., and Lowry, L.F., 1986, Sizes of walleye pollock, Theragra chalcogramma, consumed by marine mammals in the Bering Sea, Fish. Bull., 84: 192–197. Fry, M.D., and Toone, C.K., 1981, DDT induced feminization of gull embryos, Science, 213: 922–924. Garner, M.M., Lambourn, D.M., Jeffries, S.J., Hall, P.B., Rhyan, J.C., Ewalt, D.R., Polzin, L.M., and Cheville, N.F., 1997, Evidence of Brucella infection in Parafilaroides lungworms in a Pacific harbor seal (Phoca vitulina richardsi), J. Vet. Diagn. Invest., 9: 298–303. Gaskin, D.E., Holdrinet, M., and Frank., R., 1982, DDT residues in blubber of harbour porpoise, Phocoena phocoena (L)., from eastern Canadian waters during the five-year-period 1969–1973, Mammals in the Seas, FAO Fisheries Series 5, Vol. IV: 135–143. Geraci, J.R., and Lounsbury, V.J., 1993, Marine Mammals Ashore. A Field Guide for Strandings, Texas A&M Sea Grant Publications, Galveston, 305 pp. Gore, A., 1994, Introduction, in Silent Spring, Carson, R., Houghton Mifflin, Boston, MA, xv–xxvi.



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Gulland, F.M.D., 1999, Stranded seals: Important sentinels, J. Am. Vet. Med. Assoc., 214: 1191–1192. Gulland, F.M.D., Trupkiewicz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996, Metastatic carcinoma of probable transitional cell origin in free-living California sea lions (Zalophus californianus): 64 cases (1979–1994), J. Wildl. Dis., 32: 250–258. Harvell, C.D., Kim, K., Burkholder, J.M., Colwell, R.R., Epstein, P.R., Frimes, D.J., Hofmann, E.E., Lipp, E.K., Osterhaus, A.D.M.E., Overstreet, R.M., Porter, J.W., Smith, G.W., and Vasta, G.R., 1999, Emerging marine diseases—climate links and anthropogenic factors, Science, 285: 1505–1510. Holden, A.V., 1972, Monitoring organochlorine contamination of the marine environment by the analysis of residues in seals, in Marine Pollution and Seal Life, Ruivo, M. (Ed.), FAO, London, 266–272. Johnson, S.P., Nolan, S., and Gulland, F.M.D., 1998, Antimicrobial susceptibility of bacteria isolated from pinnipeds stranded in central and northern California, J. Zoo Wildl. Med., 29: 288–294. Kamolnick, T., Reddy, M., Miller, D., Curry, C., and Ridgway, S., 1994, Conditioning a bottlenose dolphin (Tursiops truncatus) for milk collection, Mar. Mammals Public Display Res., 1: 22–25. Keith, J.O., 1996, Residue analyses: How they were used to assess the hazards of contaminants to wildlife, in Environmental Contaminants in Wildlife: Interpreting Tissue Concentrations, Beyer, W.N., Heinz, G.H., and Redmon-Norwood, A.W. (Eds.), Lewis Publishers, Boca Raton, FL, 494 pp. Klamer, J.R., Laane, W.P.M., and Marquenie, J.M., 1991, Sources and fate of PCBs in the North Sea: A review of available data, Water Sci. Technol., 24: 77–85. LaPointe, J.M., Duignan, P.J., Marsh, A.E., Gulland, F.M., Barr, B.C., Naydan, D.K., King, D.P., Farman, C.A., Huntingdon, K.A.B., and Lowenstine, L.J., 1999, Meningoencephalitis due to a Sarcocystis neurona-like protozoan in Pacific harbor seals (Phoca vitulina richardsi), J. Parasitol., 84: 1184–1189. Lowry, L.F., Frost, K.J., and Loughlin, T.R., 1989, Importance of walleye pollock in the diets of marine mammals in the Gulf of Alaska and Bering Sea, and implications for fishery management, in Proceedings of the International Symposium on the Biology and Management of Walleye Pollock, University of Alaska Sea Grant Report, 701–726. Marine Mammal Commission, 1998, Marine Mammals and Persistent Ocean Contaminants. Proceedings of the Marine Mammal Commission Workshop, Keystone, CO, Oct. 12–15, 150. Martineau, D., Lagace, A., Beland, P., Higgins, R., Armstrong, D., and Shugart, L.R., 1988, Pathology of stranded beluga whales (Delphinapterus leucas) from the St. Lawrence estuary, Quebec, Canada, J. Comp. Pathol., 98: 287–311. Martineau, D., Lair, S., De Guise, S., Lipscomb, T.P., and Beland, P., 1999, Cancer in beluga whales from the St. Lawrence estuary, Quebec, Canada: A potential biomarker of environmental contamination, J. Cetacean Res. (Spec. Iss.), 1: 249–265. McCarthy, J.F., and Shugart, L.R., 1990, Biomarkers of Environmental Contamination, CRC Press, Boca Raton, FL. McKenzie, C., Rogan, E., Reid, R.J., and Wells, D.E., 1997, Concentrations and patterns of organic contaminants in Atlantic white-sided dolphins (Lagenorhynchus acutus) from Irish and Scottish coastal waters, Environ. Pollut., 98: 15–27. Measures, L.N., and Olson, M., 1999, Giardiasis in pinnipeds from eastern Canada, J. Wildl. Dis., 35: 779–782. Morris, W., 1975, The American Heritage Dictionary of the English Language, Houghton Mifflin, Boston, MA, 1550 pp. National Marine Fisheries Service, 1992, Recovery Plan for the Steller Sea Lion (Eumetopias jubatus), National Marine Fisheries Service, Silver Spring, MD, 92. National Research Council, 1991, Animals as Sentinels of Environmental Health Hazards, National Academy Press, Washington, D.C., 160 pp. National Research Council, 1996, The Bering Sea Ecosystem, National Academy Press, Washington, D.C., 320 pp. National Research Council, 2000, Marine Mammals and Low Frequency Sound, National Academy Press, Washington, D.C., 146 pp.


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O’Hara, T.M., Krahn, M.M., Boyd, D., Becker, P.R., and Philo, L.M., 1999, Organochlorine contaminant levels in Eskimo harvested bowhead whales of Arctic Alaska, J. Wildl. Dis., 35: 741–752. Olsen, M.E., Roch, P.D., Stabler, M., and Chan, W., 1997, Giardiasis in ringed seals from the western Arctic, J. Wildl. Dis., 33: 646–648. Osterhaus, A.D.M.E., and Vedder, E.J., 1988, Identification of a virus causing recent seal deaths, Nature, 335: 20. Osterhaus, A.D.M.E., Yang, H., and Spikers, H.E., 1985, The isolation and partial characterization of a highly pathogenic herpesvirus from the harbor seal (Phoca vitulina), Arch. Virol., 86: 239–251. Parveen, S.R., Murphree, L., Edmiston, L., Kaspar, C.W., Portier, K.M., and Tamplin, M.L., 1997, Association of multiple-antibiotic-resistance profiles with point and non-point sources of Escherichia coli in Apalachicola Bay, Appl. Environ. Microbiol., 63: 2607–2612. Peakall, D., 1992, Animal Biomarkers as Pollution Indicators, Chapman & Hall, London, 291. Reddy, M., Echols, S., Finklea, B., Busbee, D., Reif, J., and Ridgway, S., 1998, PCBs and chlorinated pesticides in clinically healthy Tursiops truncatus: Relationships between levels in blubber and blood, Mar. Pollut. Bull., 36: 892–903. Reddy, M.L., Reif, J.S., Bachand, A., and Ridgway, S.H., in press, Opportunities for using Navy marine mammals to explore associations between organochlorine contaminants and unfavorable effects on reproduction, Sci. Total Environ. Reijnders, P.J.H., 1986, Reproductive failure in common seals feeding on fish from polluted coastal waters, Nature, 324: 456–457. Reijnders, P.J.H., 1988, Ecotoxicological perspectives in marine mammalogy: Research principles and goals for a conservation policy, Mar. Mammal Sci., 4: 91–102. Richardson, W.J., Greene, C.R., Malme, C.I., and Thomson, D.H., 1995, Marine Mammals and Noise, Academic Press, San Diego, CA, 576 pp. Ridgway, S., 1997, Who are the whales? Bioacoustics, 8: 3–20. Ridgway, S., and Reddy, M., 1995, Residue levels of several organochlorines in Tursiops truncatus milk collected at varied stages of lactation, Mar. Pollut. Bull., 30: 609–614. Robbins, S.S., Springer, P.F., and Webster, C.G., 1951, Effects of 5-year DDT application on breeding bird population, J. Wildl. Manage., 15: 213–216. Robinson, J., 1969, Organochlorine insecticides and bird population in Britain, in Chemical Fallout: Current Research on Persistent Pesticides, Miller, M.W., and Berg, G.G. (Eds.), Charles C Thomas, Springfield, IL, 113–173. Ross, H.M., Foster, G., Reid, R.J., Jahans, K.L., and MacMillan, A.P., 1994, Brucella species infection in sea-mammals, Vet. Rec., 134: 359. Ross, P.S., 2000, Marine mammals as sentinels in ecological risk assessment, Hum. Ecol. Risk Assess., 6: 29–46. Ross, P.S., de Swart, R.L., Reijnders, P.J.H., Van Loveren, H., Vos, J.G., and Osterhaus, A.D.M.E., 1995, Contaminated-related suppression of delayed-type hypersensitivity and antibody responses in harbor seals fed herring from the Baltic Sea, Environ. Health Perspect., 103: 162–167. Ross, P.S., de Swart, R.L., Timmerman, H.H., Reijnders, P.J.H., Vos, J.G., Van Loveren, H., and Osterhaus, A.D.M.E., 1996, Suppression of natural killer cell activity in harbour seals (Phoca vitulina) fed Baltic Sea herring, Aquat. Toxicol., 34: 71–84. Schlundt, C.E., Finneran, J.J., Carder, D.A., and Ridgway, S.H., 2000, Temporary shift in masked hearing thresholds (MTTS) of bottlenose dolphins, Tursiops truncatus, and white whales, Delphinapterus leucas, after exposure to intense tones, J. Acoust. Soc. Am., 107: 3496–3508. Scholin, C.A., Gulland, F., Doucette, G.J., Benson, S., Busman, M., Chavez, F.P., Cordaro, J., DeLong, R., De Vogelaere, A., Harvey, J., Haulena, M., Lefebvre, K., Lipscomb, T., Loscutoff, S., Lowenstine, L.J., Marin III, R., Miller, P.E., McLellan, W.A., Moeller, P.D.R., Powell, C.L., Rowles, T., Silvagni, P., Silver, M., Spraker, T., Trainer, V., and Van Dolah, F.M., 2000, Mortality of sea lions along the central California coast linked to a toxic diatom bloom, Nature, 403: 80–84.



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Scott, M.D., Wells, R.S., and Irvine, A.F., 1990, A long-term study of bottlenose dolphins on the west coast of Florida, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 235–244. Smolowitz, R., and Doucette, G., 1995, The localization of saxitoxin and saxitoxin-producing bacteria in the siphons of butter clams, Saxidomus giganteus, Abstr., 26th Annual Proceedings of the International Association for Aquatic Animal Medicine, Mystic, CT, 66. Wise, S.A., Schantz, M.M., Koster, B.J., Demiralp, R., Mackey, E.A., Greenverg, T.T., Burow, M., Ostapczuk, P., and Lillestolen, T.I., 1993, Development of frozen whale blubber and liver reference materials for the measurement of organic and inorganic contaminants, Fresenius J. Anal. Chem., 345: 270–277.



2 Emerging and Resurging Diseases Debra Lee Miller, Ruth Y. Ewing, and Gregory D. Bossart

Introduction Emerging and resurging diseases affect both plants and animals worldwide. Novel zoonotic diseases usually cause concern because of their potential impacts on human health, but other diseases that can cause significant morbidity or mortality are also of concern because of their potential conservation importance. They can be especially devastating to endangered species where population levels are critically low (Harwood and Hall, 1990). For the purposes of this chapter, emerging diseases are defined as those diseases that have not been identified previously, or are considered a novel threat to the currently afflicted species (Wilson, 1999), and the chapter concentrates on diseases that have emerged in the past decade. Here resurging diseases are defined as those that historically have been documented in the species currently affected, but were considered to be eradicated or to no longer pose a significant problem. Unfortunately, it is often difficult to correctly define a disease as emerging or resurging in free-ranging wildlife. It therefore may be more appropriate to label such diseases as presumptive emerging or resurging diseases, given the paucity of historical data and the lack of baseline reference values from which to draw conclusions one way or the other. Daszak et al. (2000) describe three ways that wildlife species are exposed to emerging diseases. First, diseases emerge among wildlife species as a result of spillover from domestic species. This route has become increasingly common as domestic species encroach upon wildlife habitat, resulting in increased contact between domestic and wild animals. The introduction of canine distemper virus (CDV) to seals is a prime example of spillover to the marine environment. Initially, the etiologies of phocine morbillivirus outbreaks occurring in the 1980s were characterized serologically as phocine distemper virus (PDV) 1 and PDV-2 (Ross et al., 1992). These two strains were antigenically distinct from CDV and from each other (Visser et al., 1990). Subsequently, molecular analysis of isolates from tissues of Baikal seals (Phoca sibirica) revealed a wild-type CDV (Visser et al., 1993; Mamaev et al., 1995). Transmission of this new strain is thought to be via aerosols from domestic or feral dogs (Lyons et al., 1993). Aerosol transmission of CDV from adjacent susceptible terrestrial species such as raccoons and foxes is also possible. A very recent outbreak of CVD in Caspian seals (P. caspica) is thought to be responsible for about 10,000 deaths (Kennedy et al., 2000). The second mode of disease emergence occurs as an unfortunate consequence of efforts to restock species for conservation purposes (Daszak et al., 2000). This practice has allowed the

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translocation of hosts and pathogenic organisms, facilitating the exposure of previously naive animals to new diseases. Examples in marine mammals are currently rare, although the spread of leptospirosis was described in harbor seals (P. vitulina) during rehabilitation, probably as a result of exposure to terrestrial mammals, such as skunks (Stamper et al., 1998). The difficulty in preventing spread of disease in the open ocean environment means that, once introduced, the consequences of a novel disease could be devastating. Finally, natural phenomena, such as weather patterns like El Niño, can have profound effects on species and may greatly enhance the proliferation and/or transport of pathogenic organisms (Fauquier et al., 1998; Hoegh-Guldberg, 1999). This third mode of disease emergence is especially relevant to marine wildlife, and may be a major cause of disease resurgence (Harvell et al., 1999). Whether they are emerging or resurging, the diseases that impact marine mammals today deserve close attention, since the results are often devastating and the etiologies complex. Epizootics often involve multiple disease entities, with a primary etiology often difficult or nearly impossible to determine. For example, morbillivirus infections, which had not been documented in pinnipeds or cetaceans prior to 1988, have resulted in at least six marine mammal epizootics, and were implicated in mass mortality of the fragile Mauritanian population of Mediterranean monk seals (Monachus monachus) (Osterhaus et al., 1997; Kennedy, 1998). However, some investigators attributed the primary etiology of the monk seal mortality event to a harmful algal bloom of Alexandrium spp. (Hernández et al., 1998), resulting in considerable debate (Harwood, 1998). To solve issues such as these, multidisciplinary teams of investigators are needed. Wildlife veterinarians and biologists are now embracing the challenge of identifying disease processes occurring in wildlife species, their etiologies, and the impact they have on individuals, populations, and the species as a whole. Advanced technologies, such as the polymerase chain reaction (PCR), restriction fragment length polymorphism (RFLP), in situ hybridization, genetic sequencing, electron microscopy, and immunohistochemistry, have greatly enhanced our ability to identify disease etiologies. Similarly, advanced telemetry equipment has improved monitoring of free-ranging populations (see Chapter 38, Tagging and Tracking). Combining the laboratory-based identification of disease etiology with longterm population monitoring by field biologists is key to understanding diseases in wildlife. Given these tools, several diseases have recently been identified as either emerging or resurging in marine mammals.

Cetaceans Viral, bacterial, and neoplastic diseases are among the most important emerging and resurging diseases of cetaceans (Table 1) (also see Chapter 15, Viral Diseases; Chapter 16, Bacterial Diseases; Chapter 18, Parasitic Diseases; and Chapter 23, Noninfectious Diseases). For example, in the last decade, morbilliviruses have emerged as significant pathogens of cetaceans and pinnipeds worldwide. The origin of these viruses is undetermined, and their pathogenesis and epidemiology are just unfolding. Nucleotide sequence analysis of viral RNA isolated from Atlantic bottlenose dolphins ( Tursiops truncatus) that died in the 1987–1988 Atlantic Coast and the 1993 Gulf of Mexico epizootics indicated that the porpoise morbillivirus (PMV) and dolphin morbillivirus (DMV) are not species specific (Taubenberger et al., 1996). The 1987–1988 Atlantic Coast epizootic was a mixed infection; animals were infected with either DMV or PMV, and some animals had dual infections with both viral types. Only PMV was detected in dead animals from the 1993 Gulf of Mexico epizootic and the 1994 Irish Coast harbor porpoise (Phocoena phocoena) die-off, and


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Emerging and Resurging Diseases

TABLE 1 Identified Emerging and Resurging Diseases in Cetaceans Disease/Etiological Agent Papillomavirus

Porpoise morbillivirus

Dolphin morbillivirus

Pilot whale morbillivirus Unknown type of morbillivirus, first in baleen whale Arbovirus (Togaviridae) encephalitis Hepadnaviral hepatitis

Brucella spp.

Host Species Orcinus orca (killer whale) Tursiops truncatus (Atlantic bottlenose dolphin) Phocoena phocoena (harbor porpoise) Lagenorhynchus obscurus (dusky dolphin) Phocoena spinipinnis (Burmeister’s porpoise) Tursiops truncatus (Pacific bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Phocoena phocoena (harbor porpoise) Tursiops truncatus (Atlantic bottlenose dolphin) Stenella coeruleoalba (striped dolphin) Delphinus delphis (Pacific common dolphin) Delphinus delphis ponticus (Black Sea common dolphin) Globicephala melaena/melas (long-finned pilot whale) Balaenoptera physalus (fin whale) Orcinus orca (killer whale) Lagenorhynchus obliquidens (Pacific white-sided dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Lagenorhynchus acutus (Atlantic white-sided dolphin) Stenella coeruleoalba (striped dolphin) Delphinus delphis (common dolphin) Phocoena phocoena (harbor porpoise) Orcinus orca (killer whale) Globicephala spp. (pilot whale) Balaenoptera acutorostrata (minke whale)

Reference Bossart et al., 1997; 2000 Cassonnet et al., 1998 Van Bressem et al., 1999 Bossart and Ewing, unpublished data

Barrett et al., 1993 Taubenberger et al., 1996 Domingo et al., 1990 Lipscomb et al., 1994 Taubenberger et al., 1996 Reidarson et al., 1998; Birkun et al., 1999 Taubenberger et al., 2000 Jauniaux et al., 1998 Bossart and Ewing, unpublished data Bossart et al., 1990; Bossart, unpublished data

Foster et al., 1996 Clavareau et al., 1998 Miller et al., 1999

(Continued)


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TABLE 1 Identified Emerging and Resurging Diseases in Cetaceans (continued) Disease/Etiological Agent

Helicobacter spp. Lobomycosis Histoplasmosis Coccidioidomycosis Immunoblastic malignant lymphoma

Oral squamous cell carcinoma Renal adenoma Pulmonary carcinoma Angiomatosis

Host Species Balaenoptera physalus (fin whale) Balaenoptera borealis (sei whale) Lagenorhynchus acutus (Atlantic white-sided dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Pacific bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Stenella frontalis (Atlantic spotted dolphin) Stenella attenuata (pantropical spotted dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin) Tursiops truncatus (Atlantic bottlenose dolphin)

Reference

Fox et al., 2000 Haubold et al., 1998 Jensen et al., 1998 Reidarson et al., 1998 Bossart et al., 1997

Renner et al., 1999 Cowan and Turnbull, 1999 Ewing and MignucciGiannoni, in review Turnbull and Cowan, 1999

only DMV was recovered in the Mediterranean striped dolphin (Stenella coeruleoalba) epizootic. Taubenberger et al. (1996) proposed that cetacean morbilliviruses had actually been present in the western Atlantic prior to the European epizootics. Lipscomb et al. (1994) retrospectively examined histological specimens from the 1987–1988 Atlantic Coast epizootic for morbillivirus antigen; using immunocytochemical techniques, they detected morbillivirus antigen in 53% of the animals examined. Duignan et al. (1995a) found morbillivirus antibodies in 86% of two species of pilot whales (Globicephala melas and G. macrorhynchus) in the western Atlantic. They hypothesized that pilot whales were long-distance vectors during their trans-Atlantic migrations (Duignan et al., 1995b). Barrett et al. (1995) found that 93% of the long-finned pilot whales (G. melas) that mass-stranded between 1982 and 1993 were morbillivirus seropositive, providing further evidence that cetacean morbilliviruses are widespread, occurring in many cetacean species in the Atlantic. Interestingly, recent molecular findings of Taubenberger et al. (2000) suggest that the long-finned pilot whale is host to a different, novel type of cetacean morbillivirus, distinct from both PMV and DMV. Since the cetacean morbillivirus epizootics in Europe, the northwest Atlantic, and the Gulf of Mexico, there has been evidence of morbillivirus circulating through certain Pacific odontocete populations (Reidarson et al., 1998b; Van Bressem et al., 1998; Uchida et al., 1999). There are seropositive dusky dolphins (Lagenorhynchus obscurus), common dolphins (Delphinus delphis), and offshore bottlenose dolphins (T. truncatus) in the southeastern Pacific (Van Bressem et al., 1998). Common dolphins in the northeastern Pacific were seropositive and had viral RNA detected



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by PCR, although they did not show clinical signs of disease (Reidarson et al., 1998b). Uchida et al. (1999) reported a striped dolphin with nonpurulent meningoencephalomyelitis that stranded in Miyazaki, Japan. Using immunocytochemical techniques, they applied monoclonal anti-CDV antibodies and detected positive immunoreactivity in degenerate and intact neurons, suggesting a spontaneous morbillivirus infection. Benign mucosal and cutaneous papillomas, and/or fibropapillomas, have been characterized macroscopically and microscopically in various cetacean species. A papillomavirus etiology has been implicated for lesions in killer whales (Orcinus orca), sperm whales (Physeter macrocephalus), belugas (Delphinapterus leucas), harbor porpoises, Burmeister’s porpoises (Phocoena spinipinnis), dusky dolphins, and the offshore stock of bottlenose dolphins (Lambertsen et al., 1987; De Guise et al., 1994; Van Bressem et al., 1996; 1999; Bossart et al., 2000). Strong supportive evidence includes transmission electron microscopy (TEM), immunocytochemistry, and DNA in situ hybridization. Papillomavirus DNA was recently amplified by PCR of DNA from warts on genital slits of Burmeister’s porpoises, dusky dolphins, and bottlenose dolphins retrieved from the Peruvian coast (Cassonnet et al., 1999). Although viral diseases have had the most dramatic effects on cetaceans in the last decade, bacterial diseases are also important emerging diseases in cetaceans. Brucellosis, an apparently novel infectious disease of marine mammals with both zoonotic and economic implications, was reported in various seals, porpoises, dolphins, and a river otter (Lontra canadensis) (Foster et al., 1996), and an aborted bottlenose dolphin (Miller et al., 1999). Interestingly, retrospective studies of banked serum from stranded pinnipeds and cetaceans from the coasts of England and Wales collected between 1989 to 1995 revealed that the first positive sample occurred as early as 1990 (Jepson et al., 1997) (see Chapter 16, Bacterial Diseases). Recently, a novel Helicobacter species was cultured from the gastric mucosa of stranded Atlantic white-sided dolphins (Lagenorhyncus acutus) and identified using PCR (Fox et al., 2000). By using 16s rRNA analysis, the isolates were determined to be a novel species. By using a Warthin–Starry stain, spirochete bacteria were observed associated with proliferative lymphoplasmocytic gastritis. These findings suggest that this novel Helicobacter species may have a role in the pathogenesis of dolphin gastritis and ulceration.

Pinnipeds Toxins, neoplasia, and viral, bacterial, and parasitic diseases have all recently been identified as causing, or being associated with, significant morbidity or mortality in pinnipeds, especially in free-ranging populations (Table 2). Although the effects of morbilliviruses on pinnipeds have been dramatic, they will not be discussed further here (see Chapter 15, Viral Diseases). Domoic acid–induced morbidity and mortality may represent a resurging disease in eastern Pacific pinniped populations. Recent mortality of California sea lions (Zalophus californianus) along the central coast of California in 1998 and 2000 was attributed to harmful algal blooms (Gulland, 2000; Scholin et al., 2000). Domoic acid (DA) produced by the diatom Pseudonitzschia australis was detected in sea lion serum, urine, and feces, and in anchovy tissues (Lefebvre et al., 1999; Scholin et al., 2000). Demonstration of DA in the sea lion prey species suggests an oral route as the mode of toxin transmission. Histological examination of tissues revealed brain lesions characteristic of DA intoxication, including severe anterioventral hippocampal neuronal necrosis and marked neutrophil vacuolation within certain strata of the hippocampus and dentate gyri (Scholin et al., 2000). There have been documented cases of neurological dysfunction and mortality in sea lions, northern fur seals (Callorhinus ursinus), and dolphins (Gulland, 2000), which could have been associated with Pseudonitzschia blooms


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TABLE 2 Identified Emerging and Resurging Diseases in Pinnipeds Disease/Etiological Agent Phocine herpesvirus-1 and -2 Phocine morbillivirus

Canine distemper virus

Monk seal morbillivirus-WA Monk seal morbillivirus-G Influenza B

Coronavirus Brucella spp.

Campylobacter-like bacterium Coxiella burnetii Mycobacterium spp.

Host Species Phoca vitulina (harbor seal) Phoca vitulina (harbor seal) Pagophilus groenlandicus (harp seal) Cystophora cristata (hooded seal) Phoca hispida (ringed seal) Odobenus rosmarus rosmarus (Atlantic walrus) Halichoerus grypus (gray seal) Phoca sibirica (Baikal seal) Halichoerus grypus (gray seal) Phoca caspica (Caspian seal) Monachus monachus (Mediterranean monk seal) Monachus monachus (Mediterranean monk seal) Halichoerus grypus (gray seal) Phoca vitulina (harbor seal) Phoca vitulina (harbor seal) Phoca vitulina (harbor seal) Zalophus californianus (California sea lion) Odobenus rosmarus rosmarus (Atlantic walrus) Pagophilus groenlandicus (harp seal) Phoca hispida (ringed seal) Cystophora cristata (hooded seal) Halichoerus grypus (gray seal) Phocarctos hookeri (New Zealand sea lion) Phoca vitulina (harbor seal) Arctocephalus spp. (fur seal)

Reference Gulland et al., 1997; Harder et al., 1996 De Koeijer et al., 1998 Duignan et al., 1994; 1997 Visser et al., 1993 Kennedy et al., 1990

Mamaev et al., 1995 Visser et al., 1993 Lyons et al., 1993; Forsyth et al., 1998; Kennedy et al., 2000 Osterhaus et al., 1998 Osterhaus et al., 1998 Osterhaus et al., 2000

Bossart and Schwartz, 1990 Forbes et al., 2000 Tryland et al., 1999 Foster et al., 1996

Baker, 1999 La Pointe et al., 1999 Hunter et al., 1998


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TABLE 2 Identified Emerging and Resurging Diseases in Pinnipeds (continued) Disease/Etiological Agent

Listeria ivanovii Sarcocystis neurona-like Giardia spp.

Cryptosporidia spp. Contracaecum corderoi Ophthalmic condition

Host Species Otaria byronia (southern sea lion) Arctocephalus australis (South American fur seal) Phoca vitulina (harbor seal) Phoca vitulina (harbor seal) Phoca hispida (ringed seal) Pagophilus groenlandicus (harp seal) Phoca vitulina (harbor seal) Halichoerus grypus (gray seal) Zalophus californianus (California sea lion) Zalophus californianus (California sea lion) Monachus schauinslandi (Hawaiian monk seal)

Reference Bernardelli et al., 1996

Thornton et al., 1998 Lapointe et al., 1998 Olson et al., 1997 Measures and Olson, 1999 Deng et al., 2000

Deng et al., 2000 Fletcher et al., 1998 Banish and Gilmartin, 1992

that have occurred along the California coast over the past three decades (Walz et al., 1994). However, the DA-producing diatom P. australis did not receive much attention until a seabird mortality event occurred concurrently with a P. australis bloom in Monterey Bay, California, in 1991 (Work et al., 1993). The impacts of human and climatic activities on coastal seawater temperatures and quality may influence algal species diversity and abundance. Hernández et al. (1998) detected variable levels of numerous paralytic toxins, including decarbamoyl saxitoxin, neosaxitoxin, and gonyautoxin-1 in Mediterranean monk seal liver, kidney, skeletal muscle, and brain collected during a 1997 mortality event. The same toxins were detected in certain monk seal prey species, suggesting an available source of toxin and providing a strong indication that saxitoxins may have played a role in the monk seal mortality event. However, both the lethal toxin levels and the pharmacokinetics and baseline levels of saxitoxin in tissues of monk seals are unknown, making it difficult to interpret the toxin levels found in the animals from the 1997 epizootic (Harwood, 1998). Metastatic urogenital epithelial cell carcinomas have been reported in stranded California sea lions over the last 20 years (Gulland et al., 1996). The high prevalence of urogenital neoplasia in California sea lions suggests either a communicable infectious etiology or a common exposure to oncogenic environmental factors. Investigations of tumor etiopathogenesis have focused on the role of environmental chemical contaminants and viruses (Gulland et al., 1995; Buckles et al., 1999; Lipscomb et al., 2000). In examining cases of metastatic urogenital carcinoma, Lipscomb et al. (2000) described areas of intraepithelial neoplasia with cells containing eosinophilic intranuclear inclusion bodies. By using immunocytochemical techniques, these intranuclear inclusion bodies were shown to be positive for Epstein–Barr virus latent membrane protein. Additionally, herpesvirus-like particles were observed by TEM, and


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amplification of DNA extracted from frozen tumor samples was positive for consensus regions of herpesvirus terminase and DNA polymerase genes. Additional nucleotide sequence data indicate that the herpesvirus detected is a member of the γ-herpesvirus family. The most significant emerging bacterial disease of pinnipeds is currently brucellosis (see Chapter 16, Bacterial Diseases). Brucella spp. have been isolated from harbor seals in the eastern Pacific (Garner et al., 1997b) and from ringed (Phoca hispida) and harp seals (Pagophilus groenlandicus) near the Magdalene Islands, Gulf of St. Lawrence (Forbes et al., 2000). These marine mammal isolates are genetically distinct from currently recognized terrestrial species of Brucella and are considered novel Brucella species (Jahans et al., 1997; Bricker et al., 2000). Serological surveys for antibodies to Brucella in various species, including hooded (Cystophora cristata), harp, and ringed seals, indicate that this Brucella species is well distributed in northern Atlantic marine mammal populations (Tryland et al., 1999). Other zoonotic organisms emerging as pathogens of marine mammals are Cryptosporidium and Giardia spp. Canadian researchers investigated the prevalence of Giardia spp. and Cryptosporidium spp. in marine mammals from the Canadian western Arctic region in 1994 and 1995 and on the eastern Canadian Coast in 1997 and 1998. Giardia spp. cysts were positively detected in feces by fluorescein isothiocyanate (FITC)-labeled monoclonal antibody (Olson et al., 1997; Measures and Olson, 1999). Along the eastern coast, Giardia spp. occurred at a prevalence of 25% in gray (Halichoerus grypus) and harbor seals from the Gulf of St. Lawrence and the St. Lawrence estuary (Measures and Olson, 1999). Adult harp seals, sampled near the Magdalene Islands, Gulf of St. Lawrence, had the highest prevalence of Giardia cysts, at 50%. All pups less than 1 year of age were negative for cysts. In the western Arctic region, specifically the Holman region of the Northwest Territories, there was a 20% prevalence of Giardia in ringed seals (Olson et al., 1997). Incidentally, belugas sampled from both sites, and a northern bottlenose whale (Hyperoodon ampullatus) sampled from eastern Canada, were negative for Giardia spp. (Olson et al., 1997; Measures and Olson, 1999). Deng et al. (2000) investigated the prevalence of Cryptosporidium spp. as well as Giardia spp. in Pacific harbor seals, northern elephant seals (Mirounga angustirostris), and California sea lions from the northern California coast. They detected Cryptosporidium spp. oocysts in three California sea lions, one of which also had Giardia spp. cysts. Oocysts were then isolated and purified for PCR characterization: C. parvum and G. duodenalis were identified based on genetic characterization and morphological and immunological findings. Another protozoan, Sarcocystis spp., has been recognized as an important cause of mortality in adult Pacific harbor seals along the central California coastline (La Pointe et al., 1998). Microscopically, every case presented with marked to severe cerebellar nonsuppurative meningoencephalitis associated with S. neurona–like protozoa (La Pointe et al., 1998; Chechowitz et al., 1999). This protozoal parasite was isolated from the brain tissue from one harbor seal, and investigations are currently under way to further characterize it genetically and serologically. A helminth of emerging importance to pinnipeds is the nematode Contracaecum corderoi. From January 1992 through December 1997, C. corderoi induced gastrointestinal perforations with associated peritonitis in stranded California sea lions along the central California coast (Fletcher et al., 1998). At that time, C. corderoi had only been reported in southern fur seals (Arctocephalus australis) (Dailey and Brownell, 1972).

Manatees Currently, mortality associated with toxic algal blooms is the resurging disease with the most impact on manatees (see Chapter 22, Toxicology). From early March to late April 1996, at least 150 manatees died in an unprecedented epizootic along approximately 80 miles of the

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southwest coast of Florida (U.S. Marine Mammal Commission Annual Report to Congress, 1996). Brevetoxicosis was a primary component (Bossart et al., 1998). Grossly, severe nasopharyngeal, pulmonary, hepatic, renal, and cerebral congestion was present in all cases. Staining with interleukin-1β-converting enzyme was positive for brevetoxin in lymphocytes and macrophages in the lung, liver, and in secondary lymphoid tissues. Retrospective immunohistochemical staining of manatee tissues from an epizootic in 1982 (O’Shea, 1991) revealed widespread brevetoxin, suggesting brevetoxicosis as a component of, and the likely primary etiology for, epizootics in 1982 and 1996. As for many marine mammal species, cutaneous viral papillomatosis is an emerging disease in the Florida manatee (Trichechus manatus latirostris). Ewing et al. (1997) first reported suspected viral cutaneous papillomatosis in a captive West Indian manatee (T. manatus); diagnosis was made by light and transmission electron microscopy, which showed 45 to 50 nm spherical to hexagonal papillomavirus-like viral particles in dense arrays and smaller aggregates.

Sea Otters Parasites are emerging as a major cause of disease in the California sea otter (Enhydra lutris). Acanthocephalan parasites have long been identified as a cause of mortality in California sea otters, but in recent years the prevalence and intensity of infection appear to be increasing (Thomas and Cole, 1996). Mortality is due to peritonitis following migration of the parasites from the intestine. In a retrospective study of beached sea otters, Dailey and Mayer (1999) noted that young male otters are more frequently affected by acanthocephalans than are other animals in the population. Acanthocephalans, primarily Polymorphus spp. and Corynosoma spp., are acquired by consumption of crabs (Emerita spp. and Blepharipoda spp.) that serve as intermediate hosts for the parasites, but are not the preferred food of most otters. Dailey and Mayer (1999) hypothesize that young animals are more susceptible to infection by these parasites because of their lack of feeding experience and low social status, which leads to the foraging of less desirable food sources. Protozoans also pose a threat to sea otters. Researchers at the National Wildlife Health Center, Madison, WI, have been conducting necropsies on the threatened southern sea otter since 1992. Over the last 8 years, protozoal encephalitis was present in 8.5% of the otters received for necropsy (Thomas and Cole, 1996). Recently, Sarcocystis neurona–like protozoans and Toxoplasma gondii have been associated with encephalomyelitis and meningoencephalitis, respectively, in southern sea otters (Chechowitz et al., 1999; Rosonke et al., 1999; Cole et al., 2000). Merozoites have also been seen in skeletal muscle at multiple anatomical locations (Rosonke et al., 1999). Lindsay et al. (2000) described mostly minimal cerebral inflammation in animals examined, with only two cases showing severe fulminant meningoencephalitic sarcocystosis. They subsequently isolated protozoal merozoites from the brain of an otter with neurological disease, which were characterized as S. neurona by PCR. In general, the classic terrestrial life cycle for Sarcocystis includes an herbivore, as an intermediate host, and a carnivore or omnivore as a definitive host, but the mode of transmission to sea otters is still unclear. Another protozoan, T. gondii, has been isolated from southern sea otters, and was infective in subsequent passages through mice (Cole et al., 2000). All isolates characterized were genetically distinct, but of the same type II strain. The majority of human and pig toxoplasmosis cases are also due to the type II strain (Howe et al., 1997; Mondragon et al., 1998). It is unclear, in southern sea otters, whether the high incidence of the type II strain is due to high regional prevalence, an increased strain pathogenicity, and/or a high rate of infection. The majority of animals infected did not have severe inflammatory changes, but all presented with at least mild


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meningoencephalitis. Sea otters may be infected through ingestion of the oocyst stage, either directly from the water or by consuming filter-feeding invertebrates. Environmental contamination by feral and domestic cat populations, either directly or due to human disposal of cat feces to the municipal water supplies, might play a significant role in epidemiology of sea otter toxoplasmosis (Cole et al., 2000). Recent outbreaks of toxoplasmosis in humans resulting from inadequately treated municipal water supplies favor the latter hypothesis (Bowie et al., 1997; Isaac-Renton et al., 1998).

Polar Bears There are few novel diseases reported in polar bears (Ursus maritimus). Fatal hepatic sarcocystosis was recently reported in two polar bears from a zoo in Anchorage, Alaska (Garner et al., 1997a). The protozoa were considered to be Sarcocystis spp. based on morphology and immunohistochemistry. The point source of infection was not identified; however, fecal contamination by birds or through food fish were suspected routes. There is serological evidence that morbillivirus is endemic in the free-ranging polar bear populations of the Bering, Chukchi, and east Siberian Seas, although epidemics of disease have not been reported (Follmann et al., 1996).

Conclusion Frequency and severity of reported emerging and resurging diseases are increasing (Harvell et al., 1999). The increase may be due, in part, to improved observation and record keeping following opportunistic examinations, increased numbers of necropsies performed by pathologists rather than by biologists, and multidisciplinary investigations of recent mortality epizootics. Stranded animals, fishery by-catch, subsistence-harvested animals, and animals caught for research purposes are being more closely examined by veterinarians and pathologists. Additionally, a variety of novel technologies have enhanced identification of pathogens and toxins, so that agents may be detected in small or decomposing tissue samples. Thus, it is difficult to determine whether there is a true increase in diseases in marine mammals or merely an improvement in technology and effort. The development of long-term monitoring programs is needed to establish the significance of emerging and resurging diseases. These programs need to be transboundary, to encompass the entire migratory route of a marine mammal and the factors affecting it, and multidisciplinary. Understanding the pathogenesis of a disease, as well as its etiology and epidemiology, is paramount to understanding the potential effects of emerging and resurging diseases on a population. Accompanying the problems posed by these newly recognized infectious agents are the complications associated with the emergence of pathogen antimicrobial resistance (PAR), which has been recognized in various individual marine mammal cases (Johnson et al., 1998). Frequent use and abuse of antibiotics within both human and veterinary medicine, as well as within the agricultural industry, combined with the contamination of the environment with resistant bacteria through raw sewage spills, municipal water dumping, and agricultural and storm/flood runoff, may have important effects on marine bacteria. Care must be taken when determining the impact of resurging and emerging diseases to distinguish between diseases that were present previously but not identified and those that were truly not present. It will also be important to distinguish between primary and secondary diseases, and for secondary diseases, to determine the possible underlying causes for morbidity and mortality. Data collection and baseline life history information are key to elucidating the answers to these questions, although there are often limitations on acquiring this information, such as public indifference, limiting management policies, and inadequate funding. Regardless



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of these factors, routine and systematic sampling of animals in research, free-ranging, and captive environments must be implemented, and samples should be processed in three categories. First, samples from clinically normal animals should be analyzed to obtain normal values to use for comparisons. Second, samples from clinically normal and ill animals should be subjected to testing with currently available tests. Finally, a subsample of all collected samples should be archived for future analysis; this may prove to be the most valuable component of all. Information on disease mechanisms, pathogenesis, epidemiology, ecology, and biology can be acquired most efficiently and accurately through collaborative, international, and interdisciplinary baseline research and epizootic investigations. The authors hope that these will continue to develop, so that the role of diseases in marine mammal health and conservation can be understood.

Acknowledgments The authors thank Julia Zaias, Rosandra Manduca, Ailsa Hall, and Kirsten Gilardi for their reviews and editorial comments on this chapter, as well as all those who provided updated information on emerging and resurging diseases in marine mammals.

References Baker, A., 1999, Unusual mortality of the New Zealand sea lion Phocarctos hookeri, Auckland Islands, January–February 1998, Report of a workshop held 8–9 June 1998, Wellington, NZ, and a contingency plan for future events, New Zealand Department of Conservation, 84 pp. Banish, L.D., and Gilmartin, W.G., 1992, Pathological findings in the Hawaiian monk seal, J. Wildl. Dis., 28: 428–434. Barrett, T., Visser, I.K.G., Mamaev, L., Goatley, L., Van Bressem, M.F., and Osterhaus, A.D.M.E., 1993, Dolphin and porpoise morbilliviruses are genetically distinct from phocine distemper virus, Virology, 193: 1010–1012. Barrett, T., Blixenkrone-Moller, M., Di Guardo, G., Domingo, M., Duignan, P., Hall, A., Mamaev, A., and Osterhaus, A.D.M.E., 1995, Morbilliviruses in aquatic mammals: Report on round table discussion, Vet. Microbiol., 44: 261–265. Bernardelli, A., Bastida, R., Loureiro, J., Michelis, H., Romano, M.I., Cataldi, A., and Costa, E., 1996, Tuberculosis in sea lions and fur seals from the south western Atlantic coast, Rev. Sci. Tech. Int. Off. Epizootics, 15: 985–1005. Birkun, A., Kuiken, T., Krivokhizhin, S., Haines, D.M., Osterhaus, A.D.M.E., van de Bildt, M.W.G., Joiris, C.R., and Siebert, U., 1999, Epizootic of morbilliviral disease in common dolphins (Delphinus delphis ponticus) from the Black Sea, Vet. Rec., 144: 85–92. Bossart, G.D., and Schwartz, D., 1990, Acute necrotizing enteritis associated with suspected coronavirus infection in three harbor seals (Phoca vitulina), J. Zoo Wildl. Med., 21: 84–87. Bossart, G.D., Brawner, T.A., and Cabal, C., 1990, Hepatitis B-like infection in a Pacific white-sided dolphin (Lagenorhynchus obliquidens), J. Am. Vet. Med. Assoc., 196: 127–130. Bossart, G.D., Cray, J., Decker, S., Cornell, L.H., and Altman, N.H., 1996, Cutaneous papillomavirallike papillomatosis in a killer whale (Orcinus orca), Mar. Mammal Sci., 12: 274–281. Bossart, G.D., Ewing, R., Herron, A.J., Cray, B., Mase, B., Decker, S.J., Alexander, J.W., and Altman, N.H., 1997, Immunoblastic malignant lymphoma in dolphins: Histologic, ultrastructural, and immunohistochemical features, J. Vet. Diagn. Invest., 9: 454–458. Bossart, G.D., Baden, D.G., Ewing, R., Roberts, B., and Wright, S.D., 1998, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizootic: Gross, histologic, and immunohistochemical features, Toxicol. Pathol., 26: 276–282.


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3 Florida Manatees: Perspectives on Populations, Pain, and Protection Thomas J. O’Shea, Lynn W. Lefebvre, and Cathy A. Beck

Introduction The Florida manatee (Trichechus manatus latirostris) has been the subject of intensive research for over 25 years, using both stranding and field ecology approaches. Mandated by specific state and federal legislation, the objectives of this research have been rooted in the desire to improve manatee management for conservation of populations. Although there have been a number of different management issues that have confronted conservation efforts, the most overwhelming and persistent has been the direct mortality of manatees from accidental collisions with boats. One of the world’s most thorough and long-standing marine mammal carcass recovery and necropsy programs has clearly demonstrated that deaths of manatees from this one anthropogenic source is undisputedly a chronic, major, and growing problem (see, for example, Beck et al., 1982; O’Shea et al., 1985; Ackerman et al., 1995; Wright et al., 1995). Straightforward management solutions to this problem have been proposed, but only slowly achieved. These solutions involve a legislatively mandated policy to implement and enforce speed limits on boats in areas known to be used by manatees. To a lesser degree, solutions also involve creating sanctuaries where no boat traffic is allowed. The simple rationale is that at reduced speeds, the force of impact will be less deadly, and manatees will be more able to avoid slower boats; additionally, accidental collisions with boats cannot occur in sanctuaries where boats are excluded. Resistance to these management tools can be substantial, and some arguments against them center around incomplete knowledge of manatee population trends. However, such arguments ignore the troubling issues raised by the widespread maiming and pain inflicted on individual manatees that are struck by boats (Figure 1), escape death, and are thus not included among carcass count statistics. This overview has three related objectives. First, it provides simple documentation, descriptive summaries, and anecdotal accounts that demonstrate the extent to which maiming, and likely pain and suffering, occur in wild manatees as a result of strikes by boats. The chapter calls attention to the issues wounding raises for policy makers and managers involved with implementing boat speed zones, particularly in regard to existing laws and emerging ethical points 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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FIGURE 1 Boat-inflicted wounds on wild, living Florida manatees. (A) Multiple lacerations on dorsal tail fluke. (Photo credit: J. Reid, U.S. Geological Survey.) (B) Trunk and tail stock of adult female with completely amputated fluke. (Photo credit: T. O’Shea, U.S. Geological Survey.) (C) Lacerations of the head. (Photo credit: R. Bonde, U.S. Geological Survey.) (D) Healed severe dorsal and lateral propeller wounds. (Photo credit: K. Curtin.)

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of view. The authors suggest that considerations related to wounding should also be embraced in developing boat speed zone and sanctuary decisions, and that this issue adds a strong dimension that can override debate about manatee population trends. The strength of the science behind the latter is often misunderstood, leading to unnecessary controversy. Therefore, the second major objective is to provide a simple primer on concepts and uncertainties in manatee population biology for manatee veterinarians, rehabilitators, and biomedical specialists. Although these specialists may have little training in population ecology, they are on the front lines in manatee rescue and treatment efforts, and are often asked by the media to comment on questions related to manatee population trends. This primer is generally restricted to review of information in the published literature or widely accessible management documents. Finally, the authors submit their viewpoint that issues surrounding uncertainty in manatee population biology may be “red herrings” that detract from implementation of management actions. As humanity enters an era of growing ethical concerns for animal welfare, the degree of maiming and injury to manatees by boats will become unacceptable. Indeed, long-standing statutes that have been overdue in their application are cited to justify manatee speed zones and sanctuaries.

Maiming of Manatees in Collisions with Boats Clearly, many manatees are hit by boats, suffer pain and wounding, but survive. One of the first references to manatees being struck by boat propellers was made in the early 1940s, while by the late 1940s, biologists were using propeller scar patterns on living manatees in the wild to identify them as individuals (see historical summary in O’Shea, 1988). Although popular accounts stating that all Florida manatees bear scars from collisions with boats are not true, most carcasses examined bear scars from previous strikes (Wright et al., 1995), and a very large number of scarred manatees exist. A photoidentification system and database of scarred manatees currently maintained by the U.S. Geological Survey Sirenia Project in Gainesville, Florida (Beck and Reid, 1995) contains only individuals with distinct scars, the vast majority of which appear to have been inflicted by propeller blades or skegs (keels). This database now documents 1184 living individuals scarred from collisions with boats. Most of these manatees (1153, or 97%) have more than one scar pattern, indicating multiple strikes by boats. The severity of mutilations for some of these individuals can be astounding. These include long-term survivors with completely severed tails, major tail mutilations, and multiple disfiguring dorsal lacerations (Figures 1 and 2). These injuries not only cause gruesome wounds, but may also impact population processes by reducing calf production (and survival) in wounded females. Anecdotal observations also speak to the likely pain and repeated suffering endured by some of these individuals. For example, during fieldwork by the senior author (O’Shea) at Blue Spring and the surrounding St. Johns River, Florida, in the 1980s, known individual manatees were re-identified while snorkeling, and tracked by radiotelemetry. During snorkeling, a few individuals of known age allowed close approach, such that past scar patterns could be counted (including less-conspicuous wounds covered by gray pigmented tissue or algae). Adults with evidence of up to 19 separate hit patterns (some with multiple cuts in a single pattern) were recorded in field notes. Many individuals were struck relatively early in life (manatees can live up to 59 years) (Marmontel et al., 1996). Ages of eight individual manatees examined underwater in February 1985, and the corresponding number of strike patterns (in parentheses) by age were as follows: age 3 (12), age 3 (6), age 4 (12), age 5 (9), age 5 (11), age 6 (19), age 7 (14), and age 8 (7). In 1983, one small calf was observed with a severe dorsal mutilation trailing a decomposing piece of dermis and muscle as it continued to accompany and nurse from its mother. This individual was again severely hit in 1984, and by age 2 its dorsum was grossly


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Handbook of Marine Mammal Medicine

A

B

C

D

FIGURE 2 Underwater photographs of severe healed dorsal and tail wounds on wild, living manatees from widely separated areas in Florida. Dorsal (A) and lateral (B) mutilations of two manatees at Crystal River in northwestern peninsular Florida, where in recent decades a variety of population data suggest increasing population trends, yet severe maiming remains evident. Similar wounds (C, D) on two manatees from the southeastern Atlantic Coast, where population data do not suggest recent population increases. (Photo credits: J. Reid, U.S. Geological Survey.)



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FIGURE 3 Underwater photograph of right dorsolateral area of a 2.5-year-old wild juvenile Florida manatee struck multiple times since birth in the St. John’s River system near Blue Spring. Note the compound fracture of the rib emerging just above and to the right of the center of the photograph. Population data suggest increasing trends at this site, yet severe maiming remains evident. (Photo credit: T. O’Shea, U.S. Geological Survey.)

deformed and included a large protruding rib fragment visible in 1985 (Figure 3). While snorkeling close to this individual on January 16, 1985, patterns of 12 separate strikes by boats were counted. Despite such severe wounding, this individual remained alive in the year 2000. Carcasses examined at necropsy also often bear healed scars of multiple past strikes by boats; one extreme case, recently noted by the Florida Marine Research Institute, had evidence of more than 50 past collisions (Powell, pers. comm.). Traumatic injuries as a result of strikes by boats are also a major concern for manatee care and rehabilitation facilities (see Chapter 43, Manatees). Records maintained by the Sirenia Project since the late 1970s document rescue and rehabilitation attempts for 109 cases (69 of which died) directly linked to boat strike injuries, accounting for about 20 to 30% of the annual number of manatee rescues. The incidence of wounding by boats in Florida manatees is probably unparalleled in any marine mammal population in the world. Seals and sea lions recovered along the California coast from 1986 through 1999, for example, showed boat propeller damage in only 0.1% of 6196 live stranded individuals of six species (Goldstein et al., 1999). There is a growing sentiment in large segments of the U.S. and European public for animal welfare, animal well-being, and animal rights. One recent poll cited by Dennis (1997) found that two thirds of 1004 Americans queried by the Associated Press agreed with the statement, “An animal’s right to live free from suffering should be just as important as a person’s right to be free from suffering.” Despite modern philosophical debates on animal rights in relation to such topics as dietary use or biomedical experimentation, the inflicting of pain on animals has long been considered against most moral and ethical tenets of Western society, particularly when pain is inflicted carelessly and needlessly. Indeed, existing laws at both the state and federal levels with relevance to Florida manatees clearly reflect these tenets (Table 1), yet these laws are seldom brought to bear on the issues involving boat speed policies in Florida. The number one objective of the Florida Manatee Recovery Plan is “1. Identify and minimize causes of manatee injury and mortality” (U.S. Fish and Wildlife Service, 1996, p. 46), but the focus and debate to date has largely been on mortality only. This is due to population implications.


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TABLE 1 Florida Statutes and Federal Laws Pertaining to Injury and Wounding of Florida Manatees Florida Statutes, Title XLVI, Crimes, Chapter 828, Section 828.12 (1)

“A person who unnecessarily overloads, overdrives, torments, deprives of necessary sustenance or shelter, or unnecessarily mutilates, or kills any animal, or causes the same to be done, or carries in or upon any vehicle, or otherwise, any animal in a cruel and inhumane manner, is guilty of a misdemeanor of the first degree, punishable as provided in s. 775.082 or by a fine of not more than $5,000, or both.”

Florida Statutes, Title XXVIII, Natural Resources; Conservation, Reclamation, and Use, Chapter 370, Section 370.12 (2) (“Florida Manatee Sanctuary Act”)

“(d)…it is unlawful for any person at any time, by any means, or in any manner intentionally or negligently to annoy, molest, harass, or disturb or attempt to molest, harass, or disturb any manatee; injure or harm or attempt to injure or harm any manatee; capture or collect or attempt to capture or collect any manatee; pursue, hunt, wound, or kill or attempt to pursue, hunt, wound, or kill any manatee; …(e) Any gun, net, trap, spear, harpoon, boat of any kind … used in violation of any provision of paragraph (d) may be forfeited upon conviction.”

U.S. Marine Mammal Protection Act of 1972 (16 U.S.C. 1362, 16 U.S.C. 1372)

Sec. 3. (4) “The term ‘humane’ in the context of the taking of a marine mammal means that method of taking which involves the least possible degree of pain and suffering practicable to the mammal involved.” Sec 3. (13) “The term ‘take’ means to harass, hunt, capture, or kill, or attempt to hunt, capture, or kill any marine mammal.” Sec. 102. (a) “…it is unlawful for any person or vessel or other conveyance to take any marine mammal in waters or on lands under the jurisdiction of the United States;…”

U.S. Endangered Species Act of 1973 (16 U.S.C. 1531)

Sec. 3 (18) “The term ‘take’ means to harass, harm, pursue, hunt, shoot, wound, kill, trap, capture, or collect, or attempt to engage in any such conduct.” Sec. 9 (a) (1) “… it is unlawful for any person subject to the jurisdiction of the United States to … (B) take any such species within the United Sates or the territorial seas of the United States.”

Emphasis in italics added by authors (see also Chapter 33, Legislation).

A Primer on Manatee Population Biology: Accounting for the Confusion and Uncertainty Three related facets of Florida manatee population biology have resulted in confusing interpretations of the status of the subspecies: the estimation of population size (and thus trends in size), carcass counts (and their relationships with death and survival rates), and population modeling. These are discussed below along with their implications for manatee protection policies.

Estimation of Population Size and Trend There have been many studies in which manatee sightings from aircraft have been tallied (see summaries in Beeler and O’Shea, 1988; Ackerman, 1995). However, there are no estimates or confidence intervals for the size of the Florida manatee population that have been derived by reliable, statistically based, population-estimation techniques. This is not well understood by the public or by all individuals involved in manatee management, policy, or nonecological research programs. Nonetheless, this problem is clearly stated in the fundamental management document for the species, the Florida Manatee Recovery Plan: “Scientists have been unable to develop a useful means of estimating or monitoring trends in size of the overall manatee populations in the southeastern United States” (U.S. Fish and Wildlife Service, 1996, p. 9). In an ideal situation, biologists can determine sizes of animal or plant populations by conducting



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a census. A census is a complete count of individuals within a specified area and time period (Thompson et al., 1998). A survey, in contrast, is an incomplete count. With the exception of a few places where manatees may aggregate in clear shallow water, not all manatees can be seen from aircraft because of water turbidity, depth, surface conditions, variable times spent submerged, and other considerations. These and other factors affecting detectability of manatees in aerial surveys have been reviewed in detail by Lefebvre et al. (1995). Population estimation procedures for cetaceans and dugongs (Dugong dugon), in contrast, are based on sampling procedures that can be applied over broad, open areas. Survey techniques applied to these species allow adjustment for detectability and, thus, unlike Florida manatee surveys carried out along narrow stretches of coastline, yield unbiased estimates given certain sampling assumptions. These techniques generally involve forms of distance sampling (Buckland et al., 1993) or fixed-width transects that include methods to estimate correction factors for biases affecting detectability (Marsh, 1995). Differences between the reliability of results obtained by censuses or by sampling procedures that provide unbiased estimates, vs. simple count surveys, are often not appreciated by nonspecialists. Results obtained during typical manatee surveys yield unadjusted partial counts. These results are of value in providing information on where concentrations of manatees occur, likely relative abundance in various areas, and seasonal shifts in foci of abundance. However, the results do not provide good population estimates, nor can they reliably measure trends in populations. The counts are index values not calibrated by some known, empirically established, sampling relationship with the true numbers present. Index methods for estimating population trends in animals are flawed, because counts obtained are convolutions affected by numerous variables other than actual trends in populations—all of these variables can affect counts by altering detection probabilities in complex and unknown ways. These variables may also change with time, and their net effects on the index may not be linearly related to actual population size, obscuring the ability to understand true trends in populations. Attempts to standardize methods (e.g., air flight speed, altitude, time of day) and to adjust indices for some factors known to influence counts (e.g., temperature covariates in surveys at refugia) are important and have been followed in carrying out and interpreting results of manatee surveys. However, standardization of counting protocols does not compensate for the potentially large number of unknown or uncontrolled sources of variability in detectability (Thompson et al., 1998). Wildlife population specialists well grounded in sampling theory consider index monitoring as “an assessment protocol that collects data that usually represent at best a rough guess at population trends (and at worst may lead to an incorrect conclusion)” (Thompson et al., 1998). Thus over the years, manatee biologists have carried out numerous attempts to refine survey techniques as much as possible. These include attempts to test more sophisticated statistical approaches and to account for bias (Packard et al., 1985; 1986; Lefebvre and Kochman, 1991; Miller et al., 1998), as well as adjusting counts at aggregation sites for temperature and other covariates (Garrott et al., 1994; 1995; Ackerman, 1995; Craig et al., 1997). Nonetheless, an appropriate method for estimating the size of the entire manatee population in Florida has remained elusive. Despite these caveats, many biologists consider index approaches useful as opposed to the alternative of doing nothing (Fowler and Siniff, 1992). Thus, various aerial counts have been made in Florida since 1967, and the results from these numerous efforts have provided a longterm historical record. This large body of work (for review, see Ackerman, 1995) has led to the perception by nonspecialists that actual population size and trend are being monitored. Because it is likely that most manatees in Florida visit warm water sources, where they may occur in large numbers during periods of especially cold weather, surveys have been made at most of these places at such times each winter since the 1970s. During the initial years of such efforts, the most consistent high number obtained while circling these sites was considered a “minimum


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estimate” for numbers of manatees using that aggregation site, and the practice has been to sum these for each winter aggregation site and provide a “minimum estimate” for the size of the manatee population in Florida. These efforts did not consider manatees not counted, manatees tallied twice or more, manatees that may have moved between aggregation sites in short periods between high counts on different days, or manatees that were outside of the intensive survey areas. These “minimum estimates” are misnomers in that they are entirely different from the terminology used by population biologists for true population estimates based on sampling theory. The “minimum estimate” in 1978 was “at least 800–1000 manatees,” and in 1985 a summation of high counts made under unusually good conditions at aggregation sites was about 1200 manatees (see review by O’Shea, 1988). Confusion was further engendered when in 1990 the Florida legislature mandated “an impartial scientific benchmark census of the manatee population to be conducted annually” (Florida Statute 370.12.5a), despite recognition by scientists that a valid census was infeasible. In response, however, state resource agencies and cooperators have carried out intense synoptic surveys at simultaneous or nearly simultaneous times each year during winter. These surveys cover all known aggregation sites and most intervening areas, typically covering all areas in 1 or 2 days (Ackerman, 1995). Results of these index surveys are what are commonly, but incorrectly, cited as population estimates for Florida manatees. The first such survey in 1991 resulted in a count of 1268 manatees; a second survey 3 to 4 weeks later yielded a count of 1465. A year later the count was 1856. In January 1996, 2274 manatees were seen, and in the next month a count of 2639 was made. The most recent counts during two synoptic surveys in winter 1999–2000 were 1629, followed by 2222 10 days later. The wide variability in these numbers (differences of hundreds of animals within days or weeks, and a near doubling in 5 years) illustrates the unreliability of such counts as population estimates. This unreliability was further underscored when at least 150 manatees died during a red tide in southwestern Florida in early 1996 (Bossart et al., 1998), but the synoptic survey count for the west coast of Florida in January 1997 remained similar to that in 1996, prior to the die-off. Although over a 20- to 25-year period, counts have increased, perhaps reflecting an increase in the actual population in some of the regions surveyed over some segments of this time, the relationships between any of these numbers and the true population size remain unknown. Count data collected over multiple years from specific locations have also been analyzed for trends over time (Garrott et al., 1994; Ackerman, 1995; Craig et al., 1997). Conclusions about potential trends at specific sites may be stronger when they stem from more than one kind of data set. This can include combining inferences from counts, modeling population growth rates from survival and reproduction data (see below), examining carcass count data (see below), and weighing auxiliary information, such as habitat quality and factors promoting or reducing likelihood of survival, reproduction, or migration. This would provide a weight-of-evidence approach to aid policy makers and managers, based on a greater amount of information than count indices. Positive trends were observed in counts from the 1970s to early 1990s at Blue Spring (based on individual identification rather than aerial survey) and Crystal River, highly protected winter aggregation sites (Ackerman, 1995). Eberhardt and O’Shea (1995) showed that manatees at these two areas also had high population growth rates based on modeling of reproduction and survival data (but lower than rates of increase in counts, which were also influenced by immigration). Index counts adjusted for temperature and other covariates at several important power plant aggregation sites on the Atlantic Coast showed an increasing trend over 15 winters (ending in 1991–1992), whereas indices at one aggregation site in southwestern Florida (near Fort Myers) showed no trend (Garrott et al., 1994); previous analyses based on a 9-year period were also conducted by Garrott et al. (1995). This led to guarded speculation that manatee population trends on the Atlantic Coast may also have been



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increasing concomitant with increases in the adjusted index. However, the trend computed for adjusted counts from sites on the Atlantic Coast was too high to be compatible with the low to zero population growth estimates based on survival and reproduction data (Eberhardt and O’Shea, 1995). This seemingly conflicting information was recently clarified by a reanalysis of the counts at power plants using modifications to the statistical approach. The new analysis showed that an increasing trend in this adjusted index was only likely over the first third of the 15-winter data set, but that for the rest of the period the counts had not increased (Eberhardt et al., 1999). Craig et al. (1997) used a Bayesian approach (involving data-based hierarchial modeling to account for effects likely due to observation variables, movements among sites, and population trend) to reanalyze aerial survey data for the Atlantic Coast aggregation sites between 1982 and 1992. Although this analysis indicated possible population growth in the 1980s, it also concluded that trends leveled off or decreased during the early 1990s. Thus, unlike data for manatees at the Crystal River and Blue Spring sites, the weight of evidence from the late 1970s to early 1990s shows no suggestion of a continued increase in index counts of manatees on the Atlantic Coast or at Fort Myers (which together encompass a much larger geographic segment of the distribution than Blue Spring and Crystal River). Unfortunately, there are as yet no updated published analyses on which to base any trend conclusions for count indices in these areas for the full decade of the 1990s (although such work is in progress) and no comparable data for manatees in an extensive area encompassing the coastal Everglades.

Carcass Counts, Mortality, and Survival Each year, authorities release details on the annual total number of Florida manatee carcasses recovered and their causes of death. This provides very valuable data for management in revealing sources, locations, and times of anthropogenic mortality (those most amenable to management), as well as a wealth of pathological and anatomical biological information. Carcass counts are growing, particularly in very recent years, and collision with boats remains the major identifiable cause of death. In 1995, 184 manatees were found dead in Florida and adjacent states, with 39 killed by boats, whereas by 1999 a total of 272 carcasses were recovered, with 83 killed by boats. During the first 5 months of 2000, the number of carcasses shown to be due to boat strikes was on a record pace (see Chapter 43, Manatees). Unfortunately, these carcass counts are often misunderstood as true mortality data, in the population biologist’s sense of number of deaths per unit of population (mortality as a rate). These are not mortality rate data, because the actual population size is unknown. Furthermore, carcass counts themselves are also index values, and dividing the existing “estimates” by carcass counts to obtain death rates would result in further complex convolutions (one uncalibrated index divided by another). There is no reliable knowledge of the numbers of carcasses that go undiscovered or how discovery varies spatially, seasonally, or temporally. As the number of people using Florida’s coasts continues to grow, for example, the probability of discovery and reporting is likely to increase, as is the likelihood of human-associated death. Mortality can be computed as a rate from the distribution of ages at death, using anatomical age estimation approaches on carcasses (Marmontel et al., 1997), but this requires statistical assumptions that are not always amenable to verification. However, there have been recent advances in obtaining unbiased estimates of survival rates in manatees that utilize methods based on solid statistical inference that are completely independent of carcass counts or aerial survey index data. Mortality can also be estimated from these methods (100 − % survival = % mortality). These advances are based on sight–resight models, which ironically capitalize on scarring of living manatees as markers of individual distinctiveness (O’Shea and Langtimm, 1995; Langtimm et al., 1998). These methods have not yet been applied statewide, but efforts are


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under way to increase regional coverage. Results obtained thus far for manatees in three important regions of Florida (the Big Bend coast encompassing Crystal River, the St. John’s River encompassing Blue Spring, and the Atlantic Coast), have been compatible with regional count indices and population growth models for these areas. Survival rate estimation cannot provide instant appraisals relative to status of the population for the most recent past year because of calculation requirements. This is a drawback for media and policy makers, who may prefer more immediate data even when scientifically less valuable.

Population Models Population models employ mathematical relationships based on survival and reproduction rates to calculate population growth and trends in growth. Two sets of models of manatee population dynamics have been published. A deterministic model using classical mathematical approaches and various computational procedures with data on reproduction and survival of living, identifiable manatees suggests a maximum growth rate of about 7% per year (not including emigration or immigration) (Eberhardt and O’Shea, 1995). This maximum was based on the winter aggregation at Crystal River (an area with substantial protection), as studied from the late 1970s to early 1990s, and did not require estimates of population size. The analysis showed that the chief factor affecting potential for population growth is survival of adults. Low adult survival on the Atlantic Coast (a larger region with less protection) suggested very slow or no population growth over a similar period. This modeling shows the value of using survival and reproduction data obtained from photoidentification studies of living manatees to compute population growth rates with confidence intervals, information which can be used to infer long-term trends in the absence of reliable population size estimates. However, collection of similar data has been initiated only recently for other areas of the state (notably from Tampa Bay to the Caloosahatchee River beginning in the mid-1990s), and none is available over much of the remaining areas used by manatees in southwestern Florida. Population viability analysis (PVA) is a stochastic modeling approach, which varies potential scenarios impinging on reproduction and survival over long periods, and predicts responses in population growth. A PVA was carried out based on age-specific mortality rates computed from the age distribution of manatees found dead throughout Florida from 1979 through 1992 (Marmontel et al., 1997). This method of computing survival rests on certain assumptions that were not fully testable; yet, results point out the importance of adult survival to population persistence. Given population sizes that may reflect current abundance, the PVA showed that if adult mortality as estimated for the study period were reduced by a modest amount (e.g., from about 11 to 9%), as might be accomplished by management actions such as effective boat speed regulations, the Florida manatee population would likely remain viable for many years. Slight increases in adult mortality (a likely consequence of inadequate protection) would result in extinction over the long term. Given that the number of boats registered in Florida has increased from about 440,000 in 1975 to about 800,000 today, it is probably safe to accept the PVA-based conclusion that decreased adult survival and eventual extinction is a likely future outcome for Florida manatees, unless policies to protect them are aggressively implemented.

Uncertainties on Population Status: A Red Herring? Arguments against designation of boat speed zones to protect manatees sometimes point to uncertainties about trends in population size as reasons to delay implementation of these regulations. However, the above review shows that the basis for statewide population size “estimates” of any kind is scientifically weak and unsuitable for computing trends, and that



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the weight of evidence suggesting population increases over the last two decades is strong only for two aggregation areas. Furthermore, new population analyses, based on more recent (since 1992) information, are not yet available in the peer-reviewed literature, but these will be fundamental to management decisions that are more relevant to the contemporary situation. Thus, population-based arguments against mandated actions to reduce collisions between manatees and boats have no solid footing. The increases in boat numbers and collision-caused carcass counts suggest a continuing problem, and this is underscored by the widespread evidence of pain and mutilation. There are several additional points often missed in discussions about manatee protection that render counterarguments about manatee population trend misleading and irrelevant. First, a variety of different kinds of population dynamics information is not available for much of the state, and a weight-of-evidence approach to evaluating population trend is currently impossible for these areas. Precaution dictates a conservative policy in favor of protection, in the absence of quality data. Manatees remain listed as endangered under the U.S. Endangered Species Act and are protected by the Florida Manatee Sanctuary Act of 1978 and the U.S. Marine Mammal Protection Act of 1972 (see Chapter 33, Legislation). Indeed, when protection efforts under these mandates become effective, populations will begin making slow increases. It should be remembered that when increasing trends become apparent, they are not equivalent to population recovery, but only a signal of movement toward recovery. Failure to implement or maintain protection measures simply because trends might be increasing (a position that is unsupported by published analysis of data from most of the state) would only slow progress toward full recovery. It would be poor and purely reactive management to take actions only when unequivocal evidence of decline exists. Second, the laws mandating boat speed zones for manatee protection do not link policy implementation to manatee population trend. The Florida Manatee Sanctuary Act (Florida Statutes, Title XXVIII, Section 370.12 (2)(f)) instead states: “In order to protect manatees or sea cows from harmful collisions with motorboats or from harassment, the Fish and Wildlife Conservation Commission shall adopt rules under Chapter 120…regulating the operation and speed of motorboat traffic, only where manatee sightings are frequent and it can generally be assumed, based on available scientific information, that they inhabit these areas on a regular or continuous basis.” Thus implementation of boat speed zones is directed to protect manatees from harm, not from death only, and is aimed at areas where manatees are abundant, not necessarily at areas where populations are declining. Likewise, sanctuaries have been designated in the headwaters of the Crystal River to minimize harassment by swimmers, as well as to reduce the risk of boat–manatee collisions (O’Shea 1995; Buckingham et al., 1999). Growing concern about the effects of human harassment of manatees resulted in a “Manatee Harassment Round Table Discussion” in October 1999, sponsored by the Florida Fish and Wildlife Conservation Commission. This discussion addressed the desirability of discouraging direct physical contact between people and manatees. While all would agree that the sublethal wounding of manatees by boats represents a far higher degree of harassment than any imposed by contact with humans, the issue of boating harassment, separate from boat-caused manatee deaths, has yet to receive much attention. Finally, unlike aspects of aerial count data, the overwhelming documentation of gruesome wounding of manatees leaves no room for denial. Minimization of this injury is explicit in the Recovery Plan, several state statutes, and federal laws, and implicit in our society’s ethical and moral standards and the direction of current trends in those standards. Thus, the little that can be said with reasonable scientific certainty about manatee population size and trend may be essentially irrelevant to implementation of boat speed zones and sanctuaries, the key management tools for addressing the primary and long-standing issue facing manatee conservation and protection efforts in Florida.


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References Ackerman, B.B., 1995, Aerial surveys of manatees: A summary and progress report, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 13–33. Ackerman, B.B., Wright, S.D., Bonde, R.K., Beck, C.A., and Banowetz, D.J., 1995, Trends and patterns in mortality of manatees in Florida, 1974–1992, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 223–258. Beck, C.A., and Reid, J.P., 1995, An automated photo-identification catalog for studies of the life history of the Florida manatee, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 120–134. Beck, C.A., Bonde, R.K., and Rathbun, G.B., 1982, Analyses of propeller wounds on manatees in Florida, J. Wildl. Manage., 46: 531–535. Beeler, I.E., and O’Shea, T.J., 1988, Distribution and mortality of the West Indian manatee (Trichechus manatus) in the southeastern United States: A compilation and review of recent information, National Technical Information Service Publication PB88-207980/AS, Springfield, VA, two volumes, 613 pp. Bossart, G.D., Baden, D.G., Ewing, R.Y., Roberts, B., and Wright, S.D., 1998, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizootic: Gross, histologic, and immunohistochemical features, Toxicol. Pathol., 26: 276–282. Buckingham, C.A., Lefebvre, L.W., Schaefer, J.M., and Kochman, H.I., 1999, Manatee response to boating activity in a thermal refuge, Wildl. Soc. Bull., 27: 514–522. Buckland, S.T., Anderson, D.R., Burnham, K.P., and Laake, J.L., 1993, Distance Sampling: Estimating Abundance of Biological Populations, Chapman & Hall, London, 446 pp. Craig, B.A., Newton, M.A., Garrott, R.A., Reynolds III, J.E., and Wilcox, J.R., 1997, Analysis of aerial survey data on Florida manatee using Markov chain Monte Carlo, Biometrics, 53: 524–541. Dennis, J.U., 1997, Morally relevant differences between animals and human beings justifying the use of animals in biomedical research, J. Am. Vet. Med. Assoc., 210: 612–618. Eberhardt, L.L., and O’Shea, T.J., 1995, Integration of manatee life-history data and population modeling, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 269–279. Eberhardt, L.L., Garrott, R.A., and Becker, B.L., 1999, Using trend indices for endangered species, Mar. Mammal Sci., 15: 766–785. Fowler, C.W., and Siniff, D.B., 1992, Determining population status and the use of biological indices in the management of marine mammals, in Wildlife 2001: Populations, McCullough, D.R., and Barrett, R.H. (Eds.), Elsevier Applied Science, London, 1025–1037. Garrott, R.A., Ackerman, B.B., Cary, J.R., Heisey, D.M., Reynolds, J.E., Rose, P.M., and Wilcox, J.R., 1994, Trends in counts of Florida manatees at winter aggregation sites, J. Wildl. Manage., 58: 642–654. Garrott, R.A., Ackerman, B.B., Cary, J.R., Heisey, D.M., Reynolds, J.E., and Wilcox, J.R., 1995, Assessment of trends in sizes of manatee populations at several Florida aggregation sites, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 34–55. Goldstein, T., Johnson, S.P., Phillips, A.V., Hanni, K.D., Fauquier, D.A., and Gulland, F.M.D., 1999, Human-related injuries observed in live-stranded pinnipeds along the central California coast 1986–1998, Aquat. Mammals, 25: 43–51.



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Langtimm, C.A., O’Shea, T.J., Pradel, R., and Beck, C.A., 1998, Estimates of annual survival probabilities for adult Florida manatees (Trichechus manatus latirostris), Ecology, 79: 981–997. Lefebvre, L.W., and Kochman, H.I., 1991, An evaluation of aerial survey replicate count methodology to determine trends in manatee abundance, Wildl. Soc. Bull., 19: 289–309. Lefebvre, L.W., Ackerman, B.B., Portier, K.M., and Pollock, K.H., 1995, Aerial survey as a technique for estimating trends in manatee population size—problems and prospects, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 63–74. Marmontel, M., O’Shea, T.J., Kochman, H.I., and Humphrey, S.R., 1996, Age determination in manatees using growth-layer-group counts in bone, Mar. Mammal Sci., 54: 88. Marmontel, M., Humphrey, S.R., and O’Shea, T.J., 1997, Population viability analysis of the Florida manatee, 1976–1992, Conserv. Biol., 11: 467–481. Marsh, H., 1995, Fixed-width aerial transects for determining dugong population sizes and distribution patterns, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 56–62. Miller, K.E., Ackerman, B.B., Lefebvre, L.W., and Clifton, K.B., 1998, An evaluation of strip-transect aerial survey methods for monitoring manatee populations in Florida, Wildl. Soc. Bull., 26: 561–570. O’Shea, T.J., 1988, The past, present, and future of manatees in the southeastern United States: Realities, misunderstandings, and enigmas, in Proceedings of the Third Southeastern Nongame and Endangered Wildlife Symposium, Odom, R.R., Riddleberger, K.A., and Ozier, J.C. (Eds.), Georgia Department of Natural Resources, Social Circle, GA, 184–204. O’Shea, T.J., 1995, Waterborne recreation and the Florida manatee, in Wildlife and Recreationists: Coexistence through Management and Research, Knight, R.L. and Gutzwiller, K. (Eds.), Island Press, Washington, D.C., 297–311. O’Shea, T.J., and Langtimm, C.A., 1995, Estimation of survival of adult Florida manatees in the Crystal River, at Blue Spring, and on the Atlantic Coast, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 194–222. O’Shea, T.J., Beck, C.A., Bonde, R.K., Kochman, H.I., and Odell, D.K., 1985, An analysis of manatee mortality patterns in Florida, 1976–1981, J. Wildl. Manage., 49: 1–11. Packard, J.M., Summers, R.C., and Barnes, L.B., 1985, Variation of visibility bias during aerial surveys of manatees, J. Wildl. Manage., 49: 347–351. Packard, J.M., Siniff, D.B., and Cornell, J.A., 1986, Use of replicate counts to improve indices of trends in manatee abundance, Wildl. Soc. Bull., 14: 265–275. Thompson, W.L., White, G.C., and Gowan, C., 1998, Monitoring Vertebrate Populations, Academic Press, New York, 365 pp. U.S. Fish and Wildlife Service, 1996, Florida Manatee Recovery Plan, 2nd revision, U.S. Fish and Wildlife Service, Atlanta, GA, 160 pp. Wright, S.D., Ackerman, B.B., Bonde, R.K., Beck, C.A., and Banowetz, D.J., 1995, Analysis of watercraftrelated mortality of manatees in Florida, 1979–1991, in Population Biology of the Florida Manatee, O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), U.S. Department of Interior, National Biological Service, Washington, D.C., Information and Technology Report No. 1: 259–268.


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4 Marine Mammal Stranding Networks Frances M. D. Gulland, Leslie A. Dierauf, and Teri K. Rowles

Introduction Stranding networks are organizations that have developed to coordinate responses to stranded marine mammals. A stranded marine mammal has been defined in the United States as “Any dead marine mammal on a beach or floating nearshore; any live cetacean on a beach or in water so shallow that it is unable to free itself and resume normal activity; any live pinniped which is unable or unwilling to leave the shore because of injury or poor health” (Wilkinson, 1991). Although some causes of strandings have been identified, the majority remain enigmatic (Geraci, 1978; Geraci et al., 1999). The public concern for the welfare of stranded marine mammals, combined with the need to coordinate and maximize the information that can be obtained from these animals, are the forces behind stranding networks. This chapter describes the aims of stranding networks and reviews the history and structure of such networks worldwide.

Objectives of Stranding Networks The goal of stranding networks is to maximize specimen and data collection pertinent to the natural history, ecology, and health of stranded marine mammals and, in some areas, to provide a humane response for a stranded marine mammal (Geraci and Lounsbury, 1993). This information is important, because most of what is known about the life history and ecology of marine mammal species that are rarely observed in the wild has been learned from stranded animals (Geraci and St. Aubin, 1979; Wilkinson and Worthy, 1999). Changes in stranding numbers may also act as early warnings for issues of management importance, such as boat strike and entanglement of marine mammals (Seagers et al., 1986). Although one of the aims of stranding networks is to rehabilitate and release live stranded animals, the importance of this activity to marine mammal conservation is contentious (St. Aubin et al., 1996; Wilkinson and Worthy, 1999). It is still unclear how likely a rehabilitated and released individual is to survive, as efforts at postrelease tracking to date have focused on limited individuals because of the expense involved (see Chapter 38, Tagging and Tracking). It is also argued that the least-fit members of a population are more likely to strand, so that rehabilitating and releasing these individuals may interfere with natural selection (Wilkinson and Worthy, 1999). Furthermore, translocation of animals may enhance spread of diseases (St. Aubin et al.,

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1996; Daszak et al., 2000). To counter these arguments, examination of stranded animals during rehabilitation has allowed detection of a variety of novel infectious agents and disease processes that would have been difficult to detect in dead stranded animals, which are often too decomposed for diagnostic purposes. There is also little doubt that the general public is concerned about the welfare of live stranded marine mammals. The public attention given to animals in rehabilitation offers great opportunity for education on factors affecting marine mammal populations. In addition, some argue that there is an obligation to attempt to rehabilitate animals that strand as a result of direct anthropogenic effects, such as oil spills and entanglement in marine debris. The number of animals released after rehabilitation is usually negligible compared with the total free-living population, so the contribution to conservation by rehabilitating live stranded animals may thus be more indirect, through public exposure, involvement, and education, and through scientific research, rather than as numerical additions to wild populations. Collection of data and specimens from dead stranded animals is less controversial, but protocols still need to be established in many countries and/or regions to ensure validity of the data collected, maximum use of the information, and the willing cooperation between parties involved in a stranding network.

Stranding Networks Worldwide The degree of stranding network development varies worldwide, depending on funding availability, degree of public interest, extent of cooperation among federal, academic, and welfare organizations, facilities available, the number of strandings per year, and the duration of the existence of the network (Wilkinson and Worthy, 1999). In collecting information on stranding networks to compile this chapter, the most consistent concern of people contacted worldwide was the lack of funding. Contacts and brief descriptions of stranding networks are summarized in Table 1. A section on history is included, as developing networks may benefit from the experience of others.

TABLE 1 Examples of Stranding Networks Worldwide ARGENTINA Buenos Aires City and Province H. Castello Marine Mammal Laboratory Museo Argentino de Ciencias Naturales Avda. Angel Gallardo 470 1406 Buenos Aires E-mail: [email protected]

D. A. Albareda Acuario de Buenos Aires Avda. Las Heras 4155 Buenos Aires E-mail: [email protected]

J. Loureiro Fundación Mundo Marino Avda.X s/n Casilla de Correo n°6 7105 San Clemente del Tuyú Buenos Aires Province E-mail: [email protected]

R. Bastida and D. Rodriguez Universidad Nacional de Mar del Plata Depto de Ciencias Marinas Deán Funes 3350, 7600 Mar del Plata Buenos Aires Province E-mail: [email protected]


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Río Negro Province R. González Instituto de Biología Marina y Pesquera Alte, Storni Casilla de Correo 104 8520 San Antonio Oeste Rio Negro Fax: 54-2934-421002 E-mail: [email protected]

Chubut Province E. A. Crespo and S. N. Pedraza Marine Mammal Laboratory Centro Nacional Patagónico Blvd. Brown s/n 9120 Puerto Madryn, Chubut Fax: 54-2965-451543 E-mail: [email protected] [email protected]

Tierra del Fuego Province N. Goodall and A. Schiavini Marine Mammal Laboratory Centro Austral de Investigaciones Científicas Casilla de Correo N° 92 9410 Ushuaia Tierra del Fuego E-mail: [email protected] [email protected]

Structure Dead animals are examined and sampled for ecological studies, including age, structure, reproduction, feeding habits, genetics, virology, pollution, and parasitology. Live animals are taken to facilities (usually aquaria) for rehabilitation and monitoring of health status, where blood samples for routine health and serological tests are taken from live animals; federal and provincial laws regulate these institutions. Notes and Further Reading In Argentina there is no official stranding network, but there are several governmental and nongovernmental institutions concerned about stranding and health status of marine mammals. The Argentinean shoreline is so extensive that there are not enough groups to monitor it, but there is good communication between the research groups that work in the field. A stranding network has been in operation in Peninsula Valdéz since 1994, aimed at obtaining samples from stranded right whales; the Whale Conservation Institute collaborates with A. Carribero in this work. AUSTRALIA (Network varies by state) Queensland Michael Short Queensland Parks and Wildlife Service PO Box 2066 Cairns QLD 4870 Fax: 07-40523043 E-mail: [email protected]

Tasmania Nigel Brothers Wildlife Management Officer Kerrin Jeffrey Nature Conservation Branch GPO Box 44A Hobart, Tasmania 7001 Fax: 0362-333477 E-mail: [email protected] Antarctic Wildlife Research Unit School of Zoology University of Tasmania GPO Box 252-05 Hobart, Tasmania 7001 E-mail: [email protected]

Structure The Queensland Parks and Wildlife Service (QPWS) and the Great Barrier Reef Marine Park Authority work together to coordinate responses to strandings using the Incident Control Management System (ICMS). Most of the responses are performed by QPWS for logistical reasons. Strandings are reported on a hotline telephone number, which is diverted to a responder in the area with a mobile telephone. An e-mail listserve is used to (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) inform all network members of the status of a response. Live animals are transported to Sea World of the Gold Coast for rehabilitation. Dead animals are examined, samples banked for toxicology and genetics, and histology samples submitted to state laboratories. Jurisdiction over all marine mammals in Tasmanian waters and on the coastline falls to the Marine Unit of the Department of Primary Industries, Water and Environment (DPIWE, formerly the Parks and Wildlife Service) of Tasmania. Detailed necropsies are conducted on all cetaceans, and samples collected for morphology, pathology, toxicology, parasitology, reproductive, dietary, and aging investigations. All responses to strandings are conducted by volunteers trained to follow standard necropsy and sample collection procedures (Geraci and Lounsbury, 1993), and who are registered members of the Wildcare Organization. Samples from strandings are maintained and disseminated by the Tasmanian Museum and Art Gallery, and tracked by a database linked with that of DPIWE. History Concern over the status of dugongs initiated a formal stranding network in Queensland 3 years ago. Although dugongs remain the priority, the network now also responds to other marine mammals and turtles. The Antarctic Wildlife Research Unit (AWRU) began investigating cetacean stranding events in 1992, in response to strandings in Tasmania. The long-term goals of the unit were to gain a greater understanding of the biology and ecology of cetacean species in Tasmanian waters. It aimed to maximize the amount of scientific information collected from strandings, and build up a database of baseline data on these species. In 1996, the unit attended the first national stranding workshop coordinated by the then Australian National Parks and Wildlife Service (NPWS)—now Department of Primary Industries, Water and Environment (DPIWE)— providing protocols for the necropsy of and sample collection from stranded cetaceans. In 1998, due to the shift in priorities and goals of the NPWS, all strandings became the responsibility of the DPIWE. AWRU shifted its focus to the study of Globicephala melas, Physeter macrocephalus, and the Kogiidae, with federal funding received in 1997. Notes and Further Reading The response varies with species, dugongs being a priority, then endangered species. 90% of strandings are dead. Training courses are held regularly on the ICMS, stranding response, and sample collection. Tasmania has a relatively high number of strandings compared with other states in Australia. Although financial resources are limited, DPIWE seeks sponsorship for rescue equipment and training, and recently developed a flotation pontoon suitable for a 40-ton animal through sponsorship by the Australian Geographical Society. The Scientific Committee on Antarctic Research discourages the release of seals after being in captivity, especially to sub-Antarctic islands and the Antarctic continent. All pinniped releases must be approved by the relevant state agency, and require that a pre-release health assessment be performed. BELGIUM Administrative Coordination Management Unit of the North Sea Mathematical Models 3e en 23e Linieregimentsplein B-8400 Ostend Fax: 32-059704935 E-mail: [email protected]

Scientific Coordination University of Liege Laboratory of Oceanology Sart Tilman B6 4000 Liege Fax: 32-43663325 E-mail: [email protected] T. Jauniaux Sart Tilman B43 4000 Liege Fax: 32-43663325/4065 E-mail: [email protected]

Technical Coordination Jan Tavernier Royal Belgian Institute of Natural Sciences Rue Vautier, 29 1040 Brussels Fax: 32-026464433


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Structure Dead animals are necropsied and sampled for histopathology, parasitology, bacteriology, virology, and toxicology. The post-mortem examinations are performed according to the proceedings of the European Cetacean Society (ECS) Workshop on Cetacean Pathology (Kuiken and Hartmann, 1993) and to the proceedings of the workshop on sperm whale strandings in the North Sea (Jauniaux et al., 1999). The Marine Animals Research & Intervention Network (MARIN) also assists in marine mammal rescues. Live stranded animals are transported to rehabilitation centers (Harderwijk Delphinarium, the Netherlands for cetaceans and National Sea Life Blankenberge, Belgium for seals). History MARIN determines the cause of death of marine mammals and seabirds stranded along the Belgian coast and has performed toxicological analyses on collected samples since 1989. In 1994, MARIN expanded southward to France, in association with the “Centre de Recherche sur les Mammifères Marins,” La Rochelle. Collaboration also exists between MARIN and Naturalis, the National Museum of Natural History, Leiden, the Netherlands. Notes and Further Reading Kuiken, T., and Hartmann, M.G., 1993, Proceedings of the First European Cetacean Society Workshop on Cetacean Pathology: Dissection Techniques and Tissue Sampling, Leiden, the Netherlands, 13–14 September 1991, ECS Newsl. 17: 1–39. Jauniaux, T., Garcia Hartmann, M., and Coignoul, F., 1999, Post-mortem examination and tissue sampling of sperm whales Physeter macrocephalus, in Proceedings of Workshop: Sperm Whales Strandings in the North Sea—The Event, the Action, the Aftermath. Web sites: http://www.ulg.ac.be/fmv/anp.htm www.mumm.ac.be BRAZIL Southern Coast I. B. Moreno, P. H. Ott, and D. Danilewicz Grupo de Estudos de Mamiferos Aquaticos do Rio Grande do Sul (GEMARS) Rua Felipe Neri, 382 conj. 203 90440-150 Porto Alegre, RS Fax: 55-51267-1667 E-mail: [email protected]

Southeastern Coast Salvatore Siciliano Museo Nacional/UFRJ Dept. de Vertebrados, Setor de Mamiferos São Cristovao 20940-040 Rio de Janeiro, RJ Fax: 55-21568-1314 ext. 213 E-mail: [email protected]

Northeastern Coast Regis P. de Lima and Cristiano L. Parente Centro Mamíferos Aquáticos/IBAMA Estrada do Forte Orange, s/n° Caixa Postal 01 Ilha de Itamaracá PE 53900-000 E-mail: [email protected]

M. Cristina Pinedo Lab. Mamíferos Marinhos e Tartarugas Marinhas Dept. Oceanografia–FURG CP 474, Rio Grande–RS 96201-900 E-mail: [email protected] Also: [email protected]

J. Laílson-Brito, Jr., B. Fragoso, A. de Freitas Azevedo Universidade do Estado do Rio de Janeiro Dept. de Oceanografia Projeto MAQUA Av. São Francisco Xavier 524 sala 4018E 20550-013 Rio de Janeiro, RJ E-mail: [email protected]

Humpback Whale Project Marcia Engel Praia do Quitongo, s/n° CEP-45900-000 Caravelas, Bahia E-mail: [email protected] [email protected] http://www.criaativa. com.br/jubarte (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Marcos César de Oliveira Santos Projeto Atlantis—LABMAR Instituto de Biociências Dept. de Ecologia Geral Universidade de São Paulo Cidade Universitária São Paulo, SP E-mail: [email protected] Structure At present, there is no centralized reporting system, but there are approximately ten research groups monitoring strandings along the Brazilian coast. Stranding data are collected by separate research groups that deploy their own individual monitoring programs. Many data are collected through collaborations with media, fishermen, and the public. Although studies of marine mammals were concentrated along the south–southeastern coast, there have been recent efforts to increase efforts on the northeastern coast. Most research groups will collect stranded marine mammals, although there is no specific national legislation. Most groups are at least partially funded by research grants from the Brazilian government, but some rely only on funds from nongovernmental organizations. History Although there is no centralized database, a large proportion of the Brazilian coastline has been monitored for marine mammal strandings over the last 10 years by a number of different organizations. In some areas (south and southeast), efforts of the different groups have overlapped at some time, whereas in the north and northeast regions long stretches of coastline are not monitored. The oldest program has been maintained by Dr. M. Cristina Pinedo (FURG) since 1976 for the coast of Rio Grande do Sul state. The monitoring program surveys 120 km of beach to the north and south of the city of Rio Grande (29°20′S to 33°45′S) every 2 weeks, and the whole coastline bimonthly. The National Center for Research, Conservation and Management of Aquatic Mammals–Aquatic Mammals Center was officially created in 1998, although it had been operating previously as the “Centro Peixe-Boi” (Manatee Center) for the rehabilitation of marine manatees. Notes and Further Reading A first draft structure for a Northeastern Coast Stranding Network is under consideration by IBAMA, the Federal Environmental Agency (IBAMA/CMA Relatório No. 007-99). When effective, this network will be coordinated by the Centro Mamíferos Aquáticos/IBAMA, and operated by several organizations, including Grupo de Estudos de Cetáceos do Ceará (GECC), Centro Golfinho Rotador/Fernando de Noronha, Programa de Estudos de Animais Marinhos (PREAMAR/Bahia), and Universidade Federal do Rio Grande do Norte (UFRN/Natal). IBAMA/CMA, 1999, Relatório do primeiro workshop sobre Rede de Encalhe de Mamíferos Aquáticos do Nordeste-REMANE. IBAMA/CMA Relatório No. 007 99, 35 pp. Pizzorno, J.L.A., Laílson-Brito, J. Jr., Dorneles, P.R., Azevedo, A. de F., and Gurgel, I.M.G. do N., 1998, Review of strandings and additional information on humpback whales, Megaptera novaeangliae, in Rio de Janeiro, southeastern Brazilian coast (1981–1997), Rep. Int. Whales Comm., 48: 443–446. Lodi, L., and Barreto, A., 1998, Legal actions taken in Brazil for the conservation of cetaceans, J. Int. Wildl. Law Policy, 1: 403–411. There is a marine mammal discussion group on the Web, contactable via Drs. Laílson-Brito and B. Fragoso.


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TABLE 1 Examples of Stranding Networks Worldwide (continued) CANADA East Coast Jerry Conway Marine Mammal Advisor Department of Fisheries and Oceans P.O Box 1035, Dartmouth Nova Scotia, B2Y 4T3 E-mail: [email protected]

West Coast Ed Lochbaum Department of Fisheries and Oceans 3225 Stephenson Point Nanaimo, British Columbia V6B 5G3 E-mail: [email protected]

Structure All responses to strandings are under the auspices of, and require licensing by, the Department of Fisheries and Oceans (DFO). In Nova Scotia, strandings can be reported by calling 1(800) 668-6868. The Nova Scotia Network has focused primarily on removing stranded marine mammals from where they are found and returning them to the water, as there are no holding facilities. Post-mortem examinations are performed, and samples and skeletons obtained and stored for further research. History A volunteer group in British Columbia, The Marine Mammal Research group, has attempted to serve as a stranding network for about 15 years, but is not very active currently. The Nova Scotia Stranding Network has existed for about 8 years. It has experienced a high turnover and has encountered difficulties at times primarily because the volunteers are university students and move on. After a couple of years of relative inactivity, it is re-grouping. Notes and Further Reading St. Lawrence beluga strandings have been well studied by Dr. Martineau and co-workers (see Chapter 22, Toxicology; Chapter 23, Noninfectious Diseases). The Nova Scotia Stranding Network has been associated with the rescue and recovery work carried out by East Coast Ecosystems with the northern right whale in the Bay of Fundy. CARIBBEAN Nathalie Ward Eastern Caribbean Cetacean Network Box 5, Bequia St. Vincent and the Grenadines West Indies or P.O. Box 573 Woods Hole, MA 02543, USA Fax: 508-548-3317 E-mail: [email protected] Structure The Eastern Caribbean Cetacean Network (ECCN) is a regional, volunteer network that records sightings and strandings of marine mammals in the eastern Caribbean. The ECCN is a research affiliate of the Smithsonian (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Institute’s Marine Mammal Laboratory in Washington, D.C., and is sponsored by the United Nations Environment Program. It offers educational programs and workshops for children and adults, and training sessions for field identification and stranding protocols. Funding is provided by a number of nonprofit conservation organizations. The ECCN does not currently have a formal rescue or rehabilitation program nor a specimen collection. History The ECCN was founded in 1990 as a grassroots effort to identify whale and dolphin species of the eastern Caribbean. From 1990 to 1997, the facility was housed at the Museum of Antigua and Barbuda. As of June 1998, ECCN outreach programs have been housed in Bequia, St. Vincent and the Grenadines. The ECCN was founded by Nathalie Ward in response to the paucity of information available on cetaceans in the region. Notes and Further Reading The ECCN educational tools include a Field Guide to Whales and Dolphins of the Caribbean, available from Gecko Productions, Inc., P.O. Box 573, Woods Hole, MA 02543, U.S.A. CROATIA Dra s˘ ko Holcer Croatian Natural History Museum Department of Zoology Demetrova 1 HR-10000 Zagreb Fax: 385-1-4851644 E-mail: [email protected]

Caterina Maria Fortuna Adriatic Dolphin Project Tethys Research Institute HR-51551 Veli Lo s˘ inj E-mail: [email protected]

Structure The network includes the Ministry of Agriculture and Forestry through its connection with fishermen (primarily Fishing Inspectorate), the Ministry of Maritime Affairs through harbor masters’ offices, the Ministry of Internal Affairs through the Marine Police, and the Ministry of Defense through the National Center for Information and Alert. The ministries inform their offices of the project, and ask them to forward all information to the Croatian Natural History Museum (CNHM). Upon receipt of information on stranded animals, a team from the CNHM or the national stranding center goes to the site. Depending upon the animal’s condition, the team may collect the animal and transport it to Zagreb for post-mortem examination, or do a basic field examination, including species identification, measurements, collection of tissues and other samples (teeth, stomach contents), and determination of cause of death if possible. History In 1994, the Nature Protection Law was adopted under which a Special Act (Rule Book on Protection of Certain Mammalian Species, Mammalia) listing all protected species was issued in 1995. In this, bottlenose (Tursiops truncatus) and common dolphins (Delphinus delphis) were listed as protected species, but the Act extended legal protection to all other cetacean species that may be found in the Croatian part of the Adriatic Sea. Special Act (Rule Book on Compensation Fees for Damage Caused by Unlawful Actions on Protected Animal Species) was issued in 1996 by the same authority. Fines for deliberate killing or for actions that may cause damage or disturbance to cetaceans were set. The CNHM, in conjunction with the Adriatic Dolphin Project, tried to organize a stranding network at the national level in 1997. Notes and Further Reading In the first years, the network worked because of the enthusiasm of people involved, but lack of funding has stopped it almost entirely. Occasional reports are still forwarded to the CNHM, and depending on personal judgment, some stranded animals are collected. Information on strandings and carcasses is also occasionally collected by the veterinary faculty in Zagreb.


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TABLE 1 Examples of Stranding Networks Worldwide (continued) DENMARK Nature and Wildlife Section National Forest and Nature Agency Ålholtvej 1 DK-6840 Oksbøl Fax: 45-76541046 E-mail: [email protected]

Fisheries and Maritime Museum Tarphagevej 2 DK-6710 Esbjerg V Fax: 45-76122010 Web site: http://www.fimus.dk

Zoologisk Museum Universitetsparken 15 DK 2100 Copenhagen Ø Fax: 45-35321010 Web site: http://www.zmuc.dk

Structure Since 1993, the network has been run cooperatively by the National Forest and Nature Agency, the Fisheries and Maritime Museum in Esbjerg, and the Zoological Museum of the University of Copenhagen. Stranding events are reported either directly to the museums or through the regional forest districts. All cetacean strandings are recorded and all specimens other than harbor porpoises are examined. A standard autopsy is performed on all suitable animals. Harbor porpoises are only collected within the framework of special projects. A record of available data and specimens for research are kept by the two museums, and a special tissue bank is associated with the network. A list of samples will be made available as a read-only database on the forthcoming Web site of the network. History In 1885, upon an inquiry by the Zoological Museum, the Danish Ministry of Interior Affairs set up a notification procedure for its rescue service officers, receiver of wrecks, and other local representatives who by telegraph were to report strandings of “unusual sea animals” to the museum. Although the museum received frequent reports, the prime scope of this network was to obtain rare specimens, not to record all strandings, nor to provide the basis for analyses and management. The more common species therefore remained unrecorded. This procedure lasted until about 1980, when the Zoological Museum and the Fisheries and Maritime Museum initiated a formal stranding network, aiming to collect as much information and as many specimens as possible. This network has been improved several times since, most recently with the launching of a contingency plan in 1993, involving the forest districts of the National Forest and Nature Agency. Notes and Further Reading A comprehensive review of Danish whale strandings was published in 1995 by Kinze covering the period 1575 to 1991. The first report covering the period 1992 to 1997 was published in 1998 (Kinze et al., 1998). Kinze, C.C., 1995, Danish whale records 1575–1991 (Mammalia, Cetacea), Review of whale specimens stranded, directly or incidentally caught along the Danish coasts, Steenstrupia, 21: 155–196. Kinze, C.C., Tougaard, S., and Baagøe, H.J., 1998, Danske hvalfund i perioden 1992–1997 [Danish whale records (strandings and incidental catches) for the period 1992–1997], Flora Fauna, 104: 41–53. [In Danish with English summary.] FRANCE Centre de Recherche sur les Mammifères Marins (CRMM) Institut de la Mer et du Littoral Port des Minimes 17000 La Rochelle Fax: 33-(0)-546449945 E-mail: [email protected] (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Structure Strandings along the entire coastline are reported to authorities, which contact the local field operators authorized by the French Environment Office. These field operators are volunteers trained to respond to dead marine mammal strandings. When fresh, but dead, animals are dissected and samples collected for current or future studies (aging, stomach content analysis, ecotoxicology, genetics, reproductive biology, microbiology, parasitology, and pathology). For live stranded cetaceans, specialized personnel organize the rescue, or request euthanasia of the animal if its condition is too poor. Live stranded seals are taken to Océanopolis, Brest, or CRMM, La Rochelle, for rehabilitation. History The French stranding network was set up in 1971. All reported strandings are recorded in a database managed by the CRMM in La Rochelle. To date, over 8500 strandings have been recorded. Until 2000, administration of the network was funded mainly by the city of La Rochelle. It works thanks to the good-will, time, and funds of nonprofit organizations and authorized volunteers. Notes and Further Reading The CRMM produces annual reports on French marine mammal strandings. From 1990 to 1999, a mean of 460 cetaceans (4.5% of which were alive) and 40 seals (60% of which were alive) were recorded each year. There is a high rate of fisheries by-catch of small cetaceans, especially in winter. GERMANY Dr. Ursula Siebert Forschungs- und Technologiezentrum Westküste Hafentoern D 25761 Büsum Fax: 49-0-4834604199 E-mail: [email protected]

H. Benke Director, Deutsches Museum für Meereskunde und Fischerei Katharinenberg 14–20 D 18439 Stralsund

M. Stede Staatliches Veterinäriantersuchungsamt für Fische und Fischwaren Schleuenstrasse D 27472 Cuxhaven

Structure Live stranded seals are taken to the Seal Station Friedrichskoog, and live stranded small cetaceans to the Delfinarium Harderwijk, the Netherlands, for rehabilitation. By-caught or stranded carcasses are taken to the Westcoast Research and Technology Center, University of Kiel for examination. If transportation cannot be organized in a few hours, carcasses are stored in one of the 21 freezers distributed along the coast of the North and Baltic Seas. Post-mortem examinations are performed according to Kuiken and Hartmann (1993). Depending upon the state of preservation and findings at necropsy, samples for histology, bacteriology, virology, parasitology, serology, and toxicology may be collected. Additional investigations include age determination, reproductive biology, genetics, stomach content analysis, and skeleton archiving. History The major harbor seal die-off of 1988–1989 in northern Europe led to the development of a well-functioning stranding network for marine mammals. Notes and Further Reading The majority of strandings of marine mammals in German waters occur along the coast of Schleswig–Holstein (100 to 150 cetaceans, 350 to 450 seals per year). Kuiken, T., and Hartmann, M. G., 1993, Proceedings of the First European Cetacean Society Workshop on Cetacean Pathology: Dissection Techniques and Tissue Sampling, Leiden, the Netherlands, 13–14 September 1991, ECS Newsl., 17: 1–39.


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TABLE 1 Examples of Stranding Networks Worldwide (continued) GREECE Dr. Alexandros Frantzis Institute of Marine Biological Resources National Centre for Marine Research Agios Kosmas GR-166 04 Hellenikon Fax: 301-9811713 E-mail: [email protected] Structure Whenever port authorities are informed of a cetacean stranding in their area of responsibility, they inform the National Centre for Marine Research (NCMR) via a stranding report. However, this does not always happen, nor are the port police always aware of stranded cetaceans. Stranding reports may contain information on the place, date, time, number of animals, their total length, plus other measurements, species, sex, cause of death, comments, and possibly photographs. Due to lack of specific knowledge and experience in most cases, all information provided by nonspecialized persons is considered suspect, except the fact that a stranding did occur. When a stranding is unusual (e.g., mass strandings) or seems to have a particular value (rare cetacean species), additional information is gathered by contacting people who saw the stranded cetacean, searching for photographic documents, and/or going to the site. Reports are retained for further analysis only when accompanied by photographs that allow species identification, or when a good description is accompanied by a precise total length. History Occasional efforts to record cetacean strandings in Greece began in the late 1980s. However, the formal start of a network came at the end of 1991, when morbillivirus infection of Mediterranean striped dolphins reached the Hellenic Seas, and the increasing number of stranded animals became disturbing. The NCMR and the Hellenic Society for the Study and Protection of the Monk Seal (HSSPMS) took the initiative to inform portpolice authorities formally about the necessity of gathering stranding data and samples. A special stranding and sighting form was prepared and distributed to competent authorities all along the Greek coasts. Two years later, the HSSPMS ceased its cetological activity and a new nongovernmental organization, “Delphis” (Hellenic Cetacean Research and Conservation Society), started to receive stranding data (simultaneously with NCMR), and responded to cetacean strandings whenever possible. Some additional data were given to Greenpeace by its supporters. No formal stranding network yet exists in Greece. Notes and Further Reading Greece has the longest coastline of all the Mediterranean countries (more than 16,000 km) and almost 10,000 islands and islets, including many small uninhabited ones. Due to these particular geographic characteristics, Greek coasts (which are often inaccessible by land) are very difficult to monitor. However, the main reasons no formal and appropriate cetacean stranding network exists in Greece are lack of dedicated funds and, to a lesser degree, lack of a national coordinating authority. Even so, the incomplete stranding data gathered during the last 7 years have contributed significantly to our knowledge of cetaceans in Greece and the Mediterranean Sea. HONG KONG Coordinator: Dr. Thomas Jefferson Fax: 858-278-3473 E-mail: [email protected]

Local contacts: Samuel Hung, Mientje Torey, and Lawman Law MP 852-91990847

Contact within HKAFCD: Dick Choi E-mail: [email protected]

Structure The network is funded by the Hong Kong Government Agriculture, Fisheries and Conservation Department (AFCD) and assisted by a local oceanarium, Ocean Park Corporation, for veterinary support/expertise. (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) History Hong Kong, China SAR, formally established a cetacean stranding network in 1994, although limited data have been collected since 1973. Notes and Further Reading Parsons, E.C.M., and Jefferson, T. A., 2000, Post-mortem investigations on stranded dolphins and porpoises from Hong Kong waters, J. Wildl. Dis., 36: 342–357. ISRAEL Oz Goffman Israeli Marine Mammal Research & Assistance Center (IMMRAC) Fax: 972-52692477 E-mail: [email protected] Structure IMMRAC is in the Naval High School, in Mikhmoret, in the center of the Mediterranean coast of Israel. IMMRAC has three main interests: research, increasing public awareness, and rescue and rehabilitation. Academic support comes from the Leon Recenati Institute for Maritime Studies at the Haifa University. The rescue team consists of 30 volunteers, 3 of whom are veterinarians, and conducts simulation exercises twice a month. The personnel are divided into three teams according to the different geographic regions: north, center, and south. Necropsies are performed to establish the cause of death, with all data analyzed by Mia Roditi. IMMRAC is willing to offer assistance to neighboring countries if requested. History IMMRAC was established by a number of individuals that dedicated their free time and efforts to protecting and researching marine mammals along the coasts of Israel. Previously there had been no data on marine mammals in this region. IMMRAC conducted the first dolphin population surveys in the eastern Mediterranean, the Gulfs of Suez and Eilat, using information from trawler boats, and later from Navy vessels and diving boats. Recently, IMMRAC received, as a donation from “Tnuva,” Israel’s largest dairy producer, a research and rescue boat, which will enable daily population surveys to be performed. The IMMRAC volunteers began collecting bodies of beached dolphins in their private cars, sometimes assisted by government authorities. Notes and Further Reading IMMRAC activities led to the following findings: In 1995 Orit Barnea showed that the long snouted spinner dolphin (Stenella longirostris) lives in the Gulf of Eilat. This is the northernmost habitat for this Indian Ocean population. The rough toothed dolphin (Steno bredanensis) is found in the waters along the Israeli Mediterranean coastline, and is probably a rare but permanent resident. ITALY Marco Borri, Coordinatore Centro Studi Cetacei (CSC) Museo Zoologico “La Specola” via Romana 17 50125 Firenze Fax: 39-(0)55-225325 E-mail: [email protected]


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Structure A nationwide marine mammal stranding network is managed by the CSC of the Società Italiana di Scienze Naturali, based at the Civic Natural History Museum in Milan (Borri et al., 1997). Information on the stranding event is relayed from the stranding location, mostly by personnel from the Coast Guard, to a centralized answering service in Milan, provided at no cost by the insurance company Europe Assistance SpA. From there, the appropriate CSC correspondent from one of the 18 zones, into which the 8000 km of Italian coastline is subdivided, is alerted, and the appropriate intervention performed. CSC also coordinates research projects using samples obtained from the stranding program. History The CSC was created within the Milan Public Museum of Natural History with operational guidance from the Italian Society of Natural Sciences in 1985 at the first national conference on cetaceans in Riccione. CSC is recognized by Ministero delle Risorse Agricole, Alimentari e Forestali (Ministry of Agricultural, Food and Forest Resources) and is authorized by Ministero dell’Agricoltura e Foreste (Ministry of Agriculture and Forests) (CITES Office) and by Ministero dell’Ambiente (Ministry of Environment) (Service for the Conservation of Nature). One of the initial goals of CSC, whose aim is to unite researchers and institutions in Italy concerned with cetaceans, was to create “Progetto Spiaggiamenti” (a stranding project). This project, based upon similar projects in other countries, created a national network for the reporting and response to stranded cetaceans in 1986. In 1990, a second project was added, addressing the special needs of live stranded cetaceans. Notes and Further Reading Results of the network activities are published yearly in the Society’s proceedings (Atti della Società Italiana di Scienze Naturali). In 1986 through 1997, 2288 cetacean strandings were recorded. Of the 1724 identified species, 1054 (61.1%) were striped dolphins, 347 (20.1%) bottlenose dolphins, 99 (5.7%) sperm whales, 83 (4.8%) Risso’s dolphins, 40 (2.3%) fin whales, 40 (2.3%) long-finned pilot whales, 39 (2.3%) Cuvier’s beaked whales, with shortbeaked common dolphins, minke whales, false killer whales, and one dwarf sperm whale accounting for the remaining 1.4%. Borri, M., Cagnolaro, L., Podestà, M., and Ranieri, T., 1997, I1 Centro Studi Cetacei: dieci anni di attività (1986–1995), Natura (Milan), 88(1): 1–93. Cornaglia, E., Rebora, L., Gili, C., and Di Guardo, G., 2000, Histopathological and immunohistochemical studies on cetaceans found stranded on the coast of Italy between 1990 and 1997, J. Vet. Med., 47: 129–142. JAPAN T. K. Yamada National Science Museum 3-23-1 Hyakunin-cho Shinjuku-ku, Tokyo 164 E-mail: [email protected] Structure Local governments, aquaria, museums, research institutes, universities, and volunteers are loosely cooperating on stranding responses. The National Science Museum and Institute of Cetacean Research are responding mostly to dead strandings, the aquaria to live. There are about 100 to 200 strandings per year, of which 50 to 80 individuals are investigated to some extent. In 1999, about 50 necropsies were performed. Biological investigations, morphological research, and contaminant surveys have been conducted. (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) History The first symposium on marine mammal strandings was held in 1997 by the National Science Museum, the University of Tokyo, the Japanese Association of Zoos and Aquariums, the Institute of Cetacean Research, and the Sea of Japan Cetology Research Group. Further activities to save live strandings and to investigate dead strandings were decided upon. Training seminars have been held annually since then at the National Science Museum. Notes and Further Reading Traditionally, cetaceans have been heavily hunted for human consumption. MALDIVES H. Whitewaves Marine Research Centre Malé Republic of Maldives Fax: 960-322509/326558 E-mail: [email protected] Structure The Maldives is a country of some 1200 tiny coral islands, set upon a string of atolls, in the central Indian Ocean. Since mid-2000, an official strandings reporting scheme has been in place. Of the 1200 islands, some 200 are inhabited. Each inhabited island has a government office and government-appointed island chief. The Marine Research Centre (MRC) has sent recording forms to each island office, with instructions on how to report every marine mammal stranding. The scheme is inexpensive and is funded from the MRC budget. The main aim of the scheme is to obtain basic biological information about cetaceans in the Maldives. History Before early 2000 there was no marine mammal stranding network in the Maldives. Reports of cetacean strandings were occasionally sent to the MRC, in the capital Malé, and information on other strandings was collected by MRC staff during field trips. Notes and Further Reading Most stranded cetaceans are found floating dead at sea by fishermen. Nearly all those that wash up on islands or reefs appear to be dead at the time of stranding. There are only two known instances of live strandings to date. This, combined with the geography of the country (numerous small islands and reefs spread over a vast area of ocean, with consequent transport and communication difficulties), means that a network focusing on the welfare of live stranded marine mammals is unlikely to develop in the foreseeable future. Anderson R.C., A. Shaan, and Z. Waheed, 1999, Records of cetacean “strandings” in the Maldives, J. S. Asian Nat. Hist., 4: 187–202. MALTA Dr. A.Vella Department of Biology University of Malta Msida, MSD 06 Fax: 356-32903049 E-mail: [email protected] Structure The Director of the Environment Protection Department (EPD) is responsible for responding to strandings, and will send an inspector to the site to ensure that protocols are followed. The entities authorized to respond to a cetacean stranding are the Commissioner of Police, the Director of the Veterinary Services of Malta, field


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TABLE 1 Examples of Stranding Networks Worldwide (continued) cetacean researchers from the University of Malta, representatives of local NGOs, and the media. For dead cetacean strandings, the animal is measured, photographed, and a post-mortem examination undertaken. Legal proceedings may be undertaken if there is indication of human interaction. Specimens for further studies or for educational displays are taken to the University of Malta. For live cetacean strandings, the Director of the Veterinary Services determines the plan of action. The dolphinarium, Marineland, assists by making specialized equipment, a large treatment tank, and veterinary advice available. Fondazione Cetacea (Italy) is also willing to assist. History A cetacean stranding protocol was issued officially in March 1999, by the director of the EPD. Notes and Further Reading This protocol has been running smoothly since its establishment in March 1999. It is hoped that it will promote the proper handling of cetacean strandings. In the past, this was not the case, due to lack of available advice for inexperienced personnel. MEXICO Baja California Dr. Lorenzo Rojas-Bracho Programa Nacional de Investigación y Conservación de Mamíferos Marinos (PNICMM) c/o CICESE Ensnenada, Baja California, Tel. (6)174 50 50 al 53 ext 22115

Carribean Maria del Carmen Garcia: Parque Nacional Isla Contoy Subdirectora Tel (98) 497525 (98) 494021 Blvd Kukulkan km 4.8 ZH Cancún Q. Roo CP 77500

Gulf of Mexico Diana Madeleine AntochiwAlonzo Red de Varamientos de Yucatàn, A.C. Calle 53-E No. 232 entre 44 y 46 Fracc. Francisco de Montejo C.P. 97 200 Mérida Yucatán Tel. (9) 946 55 58 Tel./Fax. (9) 927 36 18 http://www.revay.org.mx E-mail: [email protected]

Pacific Hector Pérez-Cortés CRIP/INP Km. 1 Caretera Pichilingue – La Paz La Paz 23020 E-mail: [email protected] Structure The SOMEMMA (Mexican Society for Marine Mammalogy–Sociedad Mexicana de Mastozoologia Marina) organizes and coordinates all the groups interested in stranding response by maintaining a strandings database and assisting with obtaining permits from the National Institute of Ecology (INE) and Procuraduria Federal de Proteccion al Ambiente (PROFEPA). In Ensenada, Baja California, a new way of organizing stranding response efforts is being attempted. All people interested in strandings in the Ensenada–Tijuana corridor (NGOs, university, research institutes, individuals, and INE) were contacted, and representatives met with PROFEPA. Delegates created the subcommittee for strandings attention, an organization with government representation. (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Quintana Roo is the state that faces the Caribbean Sea, where the first stranding network on the east coast of Mexico was established in 1987. This group has concentrated mainly on manatees. History For over a decade, research groups have responded to marine mammal strandings, mainly in the southern state of Baja California Sur, where there is the highest density of marine mammalogists. Initially, each researcher worked independently, but efforts to coordinate responses are developing. A few years ago, in the northern State of Baja California, a group of students and researchers formed an NGO that focuses on marine mammal strandings, primarily California sea lions, in the Ensenada–Tijuana area. In the mid-1990s, the Attorneys General Office for the Environment (PROFEPA) was created, with almost every state in Mexico having a PROFEPA office. PROFEPA addresses any issue that affects the environment. It does not respond to strandings, but to be able to attend strandings, one must have its authorization and permits from the INE. Both PROFEPA and INE have created a number of subcommittees consisting of members of local communities to address environmental issues, from illegal fishing to pollution. Notes and Further Reading No government funding for these efforts exists, nor is there any possibility of financial support in the foreseeable future. Except for the states of Campeche and Tamaulipas, NGOs are currently attending strandings on the coasts of Veracruz, Tabasco, and Yucatán. Most of these groups formed in the last 3 to 4 years. Students mostly constitute these groups. Funding is extremely low and comes from contributions by the members. Some receive in-kind support from their local universities and aquaria. More recently, a national stranding e-mail correspondence group was created to discuss strategies and to exchange experiences. This information was kindly provided by SOMEMMA. THE NETHERLANDS Dr. Chris Smeenk National Museum of Natural History P.O. Box 9517 2300 RA Leiden Fax: 31-1-5687666 E-mail: [email protected] Structure The stranding network involves many official authorities and volunteers. It is coordinated by the National Museum of Natural History, Leiden. Stranding records are published in Lutra, the journal of the Dutch Mammal Society (Smeenk, 1995). Dead cetaceans are collected by or for the museum; most of them are frozen. A post-mortem on all suitable animals is carried out by a team of veterinarians and zoologists. Standard samples are taken for histopathology, bacteriology, virology, life-history, toxicology, and dietary studies (Kuiken and Hartmann, 1993). Live stranded animals are taken to the Marine Mammal Park at Harderwijk and to Zeehondencreche Pieterbuen. History Data and material from stranded cetaceans have been collected since about 1914. Archives and databases of strandings are kept in the National Museum of Natural History, Leiden. For some large species, records date back to the 16th century (Smeenk, 1997). Skeletal material and samples are deposited in the Leiden museum; other important osteological collections are in the Zoological Museum of Amsterdam University and in the Natural History Museum in Rotterdam.


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TABLE 1 Examples of Stranding Networks Worldwide (continued) Notes and Further Reading Addink, M.J., and Smeenk, C., 1999, The harbour porpoise Phocoena phocoena in Dutch coastal waters: Analysis of stranding records for the period 1920–1994, Lutra, 41: 55–80. Kuiken, T., and Hartmann, M.G., 1993, Proceedings of the First European Cetacean Society Workshop on Cetacean Pathology: Dissection Techniques and Tissue Sampling, Leiden, the Netherlands, 13–14 September 1991, ECS Newsl., 17: 1–39. Smeenk, C., 1995, Strandingen van Cetacea op de Nederlandse kust in 1990, 1991 en 1992, Lutra, 38: 90–104. Smeenk, C., 1997, Strandings of sperm whales Physeter macrocephalus in the North Sea: History and patterns, Bull. Inst. R. Sci. Nat. Belg. Biol., 67 Suppl.: 15–28. NEW ZEALAND Coordinator Anton van Helden Marine Mammals Collection Manager Museum of New Zealand Te Papa Tongarewa P.O. Box 467, Wellington Fax: 06443817310 E-mail: [email protected]

Pathologist Pádraig Duignan New Zealand Wildlife Health Centre I.V.A.B.S. Massey University Palmerston North Fax: 006463505714 E-mail: [email protected]

Department of Conservation Rob Suisted 58 Tory Street, Wellington E-mail: [email protected]

Genetics Dr. Scott Baker School of Biological Sciences University of Auckland Auckland E-mail: [email protected]

Volunteer Groups Project Jonah P.O. Box 8376 Symonds Street Auckland Fax: 064-95215425

Marine Watch Jim Lilley 59 Clydesdale St Linwood, Christchurch

Structure The Department of Conservation (DOC) administers the Marine Mammal Protection Act of 1978, which provides for the conservation, protection, and management of marine mammals. Among other roles, DOC is responsible for dealing with beached and stranded cetaceans and pinnipeds. Cetaceans that can be refloated are saved with the help of volunteer groups. Those that die are examined by a pathologist to determine cause of death. Samples are archived at Massey University for diagnostic tests, toxicology, and genetics. The marine mammals collection manager at the Museum of New Zealand Te Papa Tongarewa maintains a database of all cetacean strandings as well as collecting, storing, and maintaining an extensive skeletal collection. A database of cetacean genetics is maintained at the University of Auckland. History The New Zealand Stranding Network was established as a collaboration among the Museum of New Zealand, the Department of Conservation, universities, and Maori interest groups. Notes and Further Reading New Zealand has a large number of cetacean strandings with an average of 80 incidents per year representing as many as 38 species (an average of 19 species each year). In addition, stranded pinnipeds include New Zealand fur seals, subantarctic fur seals, leopard seals, and, less commonly, New Zealand sea lions and southern elephant seals, with historic records of crabeater seals. Web: http://www.massey.ac.nz Web: http://www.doc.govt.nz (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) PERU CEPEC Department of Veterinary Research Jorge Chavez 302, Pucusana Lima 20 E-mail: [email protected]

Centro Peruano de Estudios Cetologicos (CEPEC) Museo de la Fauna Marina Jorge Chavez 101, Pucusana Lima 20 E-mail: [email protected]

Structure No official marine mammal stranding network exists in Peru, but specimens are collected opportunistically by a variety of individuals and institutions, including CEPEC. Fresh or live cetacean strandings typically are utilized by locals. History CEPEC is a private institute founded in 1985 for research on the distribution, biology, pathology, and management issues of cetaceans in developing countries, with particular emphasis on the Southeast Pacific. SPAIN Valencia Region Fax: 34-963864372 E-mail: [email protected]

Murcia Region Tel: 34-968526817 and 34-689788515

Catalonia Region Fax: 34-937525710 E-mail: [email protected]

Andalusia Region Fax: 34-952229287 E-mail: [email protected]

Balearic Islands Tel: 34-971675125

Galician Region Cemma Tel./Fax: 34-981360804 E-mail: [email protected]

Euskadi Region Ambar E-mail: [email protected]

Cantabria Region Fax: 34-942281068

Canary Islands M. Andre Fax: 34-928451141 E-mail: [email protected]

Asturias Region Cepesma E-mail: [email protected] Structure Each coastal regional government, of which there are five in the Mediterranean, four in the Atlantic, and one in the Canary Islands, has a coordinator. Coordinators collaborate with the Spanish Cetacean Society, funded by the Spanish Ministry of Environment, to establish standard protocols and methods for sightings, strandings, and rehabilitation of cetaceans and sea turtles in Spanish waters. In the Canary Islands, there is no official stranding network, but the veterinary school (Marine Mammal Conservation Research Unit, Veterinary School, University of Las Palmas de Gran Canaria) has responded to 85% of cetacean strandings in the Canary Islands. There are no pinniped strandings. Once a year, a complete report on all island strandings is sent to the government of each island.


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TABLE 1 Examples of Stranding Networks Worldwide (continued) SWEDEN Mats Olsson Swedish Museum of Natural History Contaminant Research Group Box 50007 SE 104 05 Stockholm Fax: 46-8 5195 4256 E-mail: [email protected] Structure Seals found dead in fishing gear or stranded within the Baltic have been sent to the Swedish Museum of Natural History in Stockholm. Collection is by the public, the police, and the Swedish Coast Guard. The animals are examined to determine cause of death or health status. The health studies are part of the Swedish Environmental Monitoring Program run by the Swedish Environmental Protection Agency (EPA). Simultaneous annual censuses of the three seal populations are carried out by the Swedish Museum of Natural History, also funded by the Swedish EPA. History The Swedish program for stranded seals has existed since the 1970s. Notes and Further Reading Olsson, M., Andersson, Ö., Bergman, Å., Blomkvist, G., Frank, A., and Rappe, C., 1992, Contaminants and diseases in seals from Swedish waters, Ambio, 21: 561–562. UKRAINE (and Bulgaria and Georgia) Dr. Alexei Birkun E-mail: [email protected] Structure This network that includes three countries is coordinated by the BREMA laboratory in Simferopol, Crimea, and includes 6 specialists and 30 to 40 volunteers (students, school children, fishermen, officers of the Ukrainian Fish Protection Service, coastal border guards). There is no financial support for the network at present. History A cetacean stranding network has been working in the Crimea (Ukraine, Black Sea region) since 1989. In 1997, the network was extended into Bulgaria and Georgia. Notes and Further Reading Birkun, A., Jr., Stanenis, A., and Tomakhin, M., 1994, Action plan for rescue, rehabilitation and reintroduction of wild sick and traumatized Black Sea cetaceans. European research on cetaceans, 8, in Proc. 8th Annual Conf. Eur. Cetacean Soc., Montpellier, France, 2–5 March 1994, Lugano, 237 pp. Krivokhizhin, S.V., and Birkun, A.A., 1999, Strandings of cetaceans along the coasts of the Crimean peninsula in 1989–1996, European research on cetaceans, 12, in Proc. 12th Annual Conf. Eur. Cetacean Soc., Monaco, 20–24 January 1998, European Cetacean Society, Valencia, 59–62. Birkun, A., Jr., Kuiken, T., Krivokhizhin, S., Haines, D.M., Osterhaus, A.D.M.E., van de Bildt, M.W., Joiris, C.R., and Siebert, U., 1999, Epizootic of morbilliviral disease in common dolphins (Delphinus delphis ponticus) from the Black Sea, Vet. Rec., 144: 85–92. (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) UNITED KINGDOM Institute of Zoology Regent’s Park London, NW1 4RY Fax: 0207 586 1457 E-mail: [email protected]

The Natural History Museum Cromwell Road London, SW7 5BD Fax: 020 7942 5433

Wildlife Unit SAC Veterinary Science Division (Inverness) Drummondhill Stratherrick Road Inverness, IV2 4JZ Fax: 1463-711103 E-mail: [email protected]

British Divers Marine Life Rescue 39 Ingham Road, Gillingham Kent, ME7 1SB Tel./Fax: 01634-281680 E-mail: 101375,[email protected]

RSPCA Headquarters Wildlife Department Causeway Horsham West Sussex RH12 1HG http://www.rspca.org.uk

Scottish SPCA 603 Queensferry Road Edinburgh, EH4 6EA Fax: 0131 339 4777

Structure Coordination of pathological investigations of strandings in England and Wales has been conducted by the Institute of Zoology (Zoological Society of London) in collaboration with the Natural History Museum, London, since 1990. The Scottish Agricultural College Inverness has coordinated all strandings research investigations within Scotland since 1992. Post-mortem examinations are performed according to Kuiken and Hartmann (1993). Live strandings are reported to the Royal Society for the Protection of Animals (RSPCA) in England and Wales (24-hour hotline: 0870 5555999). In Scotland, the Scottish Society for the Protection of Animals (SSPCA) has several local emergency phone numbers. Inspectors from both organizations routinely attend such events. Live seals are taken to seal rehabilitation centers throughout the U.K., when deemed necessary. Live stranded cetaceans are typically attended by veterinarians, members of British Divers Marine Life Rescue (BDMLR), and other rescue groups who have an extensive network of trained volunteers throughout the U.K. There are currently no appropriate facilities for cetacean rehabilitation within the U.K. History Since 1913, the Natural History Museum in London has collected data on cetacean strandings within the U.K. In 1990, 2 years after a major epizootic of phocine distemper occurred in harbor seals in northern Europe, the U.K. Department of the Environment decided to partially fund a systematic and collaborative program of marine mammal strandings research within the U.K. This research is currently ongoing. The main goals of this new strandings research, apart from investigating any future marine mammal mass mortalities, were systematically to investigate the diseases, causes of death, and potential relationships between exposure to contaminants and health status in marine mammals in U.K. waters. A centralized U.K. database for pathological and other data resulting from the strandings projects and national marine mammal tissue archives were also established. Although originally established to investigate both cetacean and pinniped strandings in U.K. waters, the U.K. strandings program has been heavily biased toward cetaceans in recent years to comply with a number of international cetacean conservation agreements to which the U.K. is a signatory. Notes and Further Reading Approximately 200 cetaceans (mainly harbor porpoises and common dolphins) and 300 pinnipeds (mainly gray seals and common seals) typically strand within the U.K. each year. A number of key collaborating organizations, such as the Veterinary Investigation Unit, Truro, the Centre for Environment, Fisheries and Aquaculture Science; Sea Mammal Research Unit; University College Cork, Ireland; University of Aberdeen; Institute of Animal Health, Pirbright; and the Natural History Museum of Scotland, are involved in many aspects of the strandings research.


65


TABLE 1 Examples of Stranding Networks Worldwide (continued) Kuiken, T., and Hartmann, M.G., 1993, Proceedings of the First European Cetacean Society Workshop on Cetacean Pathology: Dissection Techniques and Tissue Sampling, Leiden, the Netherlands, 13–14 September 1991, ECS Newsl., 17: 1–39. UNITED STATES OF AMERICA http: //www.nmfs.noaa.gov/prot_res/PR2/Health_am_stranding_Response_program/mmhsrp.html Cetaceans, Seals, Sea Lions, Sea Turtles: Alaska NMFS Alaska Region P.O. Box 21668 Juneau, AK 99802-1668 Tel: (907) 586-7235 Fax: (907) 586-7249

Washington and Oregon NMFS Northwest Region 7600 Sand Point Way, N.E. Bldg. 1 Seattle, WA 98115-0070 Tel: (206) 526-6733 Fax: (206) 526-6736

Maine to Virginia NMFS Northeast Region One Blackburn Drive Gloucester, MA 01930-2298 Tel: (508) 495-2090

North Carolina to Texas, Puerto Rico, U.S. Virgin Islands NMFS Southeast Region 9721 Executive Center Drive St. Petersburg, FL 33716 Tel: (305) 361-4586

Sea Otters: U.S. Fish and Wildlife Service 2493 Portola Road, Suite B Ventura, CA 93003 Tel: (805) 644-1766

Manatees: Endangered Species Division U.S. Fish and Wildlife Service 75 Spring Street, S.W. Atlanta, GA 30303 Tel: (404) 679-7096

California and Hawaii NMFS Southwest Region 501 West Ocean Boulevard Suite 4200 Long Beach, CA 90802 Tel: (562) 980-4017

Polar Bears, Walrus, Sea Otters in Alaska: U.S. Fish and Wildlife Service 1011 East Tudor Road Anchorage, AK 99503-6199 Tel: (907) 786-3800

Structure Jurisdiction over cetaceans and seals and sea lions falls to the National Marine Fisheries Service (NMFS), while the U.S. Fish and Wildlife Service has jurisdiction over walrus, sea otters, and polar bears. The National Stranding Network is divided into five regions: Northwest, Southwest, Northeast, Southeast, and Alaska. Although officially part of the Southwest Region, all stranding responses in Hawaii are coordinated by the NMFS Pacific Area Protected Species Program Coordinator. Network members consist of a wide range of organizations and individuals, including government agencies, academic institutions, research institutions, rehabilitation facilities, aquaria, and interested individuals. Activities of members are coordinated by the NMFS regional coordinator. Training is available for network volunteers, primarily through a field guide (Geraci and Lounsbury, 1993), but also through newsletters and workshops. All participants are required to submit monthly stranding reports to their regional offices on which Level A, B, and C data are recorded. Level A data are minimum data to be collected at any stranding event and reported to the national office (exact location, date, initial species identification, number of animals involved, sex, length, evidence of human interaction, and condition of the animals). Level B data are basic life-history and specific event data (weather, carcass orientation, animals and human activities in area, collection of parts for age determination). Level C data are results of careful internal and external examination of animals involved, including specimen collection and preservation (Geraci and Lounsbury, 1993). Members do not receive direct funding from NMFS for stranding responses, except under special circumstances. History In 1972, the increased federal protection of marine mammals resulting from the passage of the Marine Mammal Protection Act (MMPA), combined with increased public awareness and compassion for marine mammals, highlighted a need for an organized response to marine mammal strandings beyond the Smithsonian Institution’s list of strandings. In 1977, the first Marine Mammal Stranding Workshop was held. The shortterm goals established at this workshop were to provide for a national network coordinator; to establish and (Continued)


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TABLE 1 Examples of Stranding Networks Worldwide (continued) evaluate regional reporting and notification systems; to establish standard protocols for euthanasia, transport, release, specimen requests, and disposal of stranded marine mammals; to describe clearly and periodically evaluate data collection; and to develop and maintain up-to-date inventories of all interested parties and network-authorized institutions. The long-term goals of this workshop were to develop procedures that would minimize possible threats to human health, minimize pain and suffering of live stranded animals, derive maximum scientific and educational benefits, and result in collection of normal baseline data. In 1981, regional offices and methods for network participation and reporting were established. By 1987, there was sufficient new information from strandings and enough need to standardize collection protocols that a second Marine Mammal Stranding Workshop was held. In 1991, a national stranding coordinator was appointed to define national stranding policy, standardize network operations, and enhance and support capabilities of network members. In 1992, the stranding networks were recognized within the MMPA with the addition of Title IV, the Marine Mammal Health and Stranding Response Act (Public Law 102–687). Notes and Further Reading If an unusual increase in stranding numbers occurs, a protocol for response described by Wilkinson (1996) occurs (see Chapter 5, Unusual Mortality Events). An interagency National Marine Mammal Tissue Bank and Quality Assurance Program held at the National Institute of Standards and Technology in Gaithersburg, MD was established to collect and archive tissues from marine mammals that can be used for retrospective analysis of contaminant levels. Geraci, J.R., and Lounsbury, V., 1993, Marine Mammals Ashore: A Field Guide for Stranding, Texas A&M University Sea Grant College Program, Galveston, 305 pp. St. Aubin, D.J., Geraci, J.R., and Lounsbury, V.J., 1996, Rescue, rehabilitation and release of marine mammals: An analysis of current views and practices, Proceedings of a workshop held in Des Plaines, Illinois, 3–5 December 1991, NOAA Technical Memorandum, NMFS-OPR-8, 65 pp. Wilkinson, D., and Worthy, G., 1999, Marine mammal stranding networks, in Conservation and Management of Marine Mammals, Twiss, J.R., and Reeves, R.R. (Eds.), Smithsonian Institution Press, Washington, D.C., 396–411. Wilkinson, D.M., 1991, Report to Assistant Administrator for Fisheries: Program review of the marine mammal strandings networks, U.S. Department of Commerce, NOAA, National Marine Fisheries Service, Silver Spring, MD, 171 pp. Wilkinson, D.M., 1996, National Contingency Plan for Response to Unusual Marine Mammal Mortality Events, Technical Memorandum NMFS-OPR-9, U.S. Department of Commerce, NOAA, NMFS, Silver Spring, MD, 118 pp.

Acknowledgments The authors thank K. Acevedo, M. Addink, D. Albareda, M. Andre, A. Barreto, J. Barnett, A. Birkun, M. Borri, N. Brothers, J. Conway, E. A. Crespo, E. Degollada, P. Duignan, K. Evans, D. Holcer, A. Frantzis, O. Goffman, T. Jauniaux, T. Jefferson, K. Jeffrey, P. Jepson, R. Kinoshita, C. Kinze, N. LeBoeuf, G. Notabartollo di Sciara, M. Olsson, E. Poncelet, J. A. Raga, B. Reid, L. Rojas, K. Rose, V. Ruoppolo, M. Short, S. Siciliano, U. Siebert, C. Smeenk, K. Soto, K. Van Waerebeek, N. Ward, A.Vella, and T. Yamada, for providing information on stranding networks, and Ailsa Hall for reviewing this chapter.

References Daszak, P., Cunningham, A.A., and Hyatt, A.D., 2000, Emerging infectious diseases of wildlife—Threats to biodiversity and human health, Science, 287: 443–449. Geraci, J.R., 1978, The enigma of marine mammal strandings, Oceanus, 21: 38–47.



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Geraci, J.R., and Lounsbury, V., 1993, Marine Mammals Ashore: A Field Guide for Strandings, Texas A&M University Sea Grant College Program, Galveston, 305 pp. Geraci, J.R., and St. Aubin, D.J., 1979, Biology of marine mammals: Insights through strandings, Final Report MMC-77/13 to the U.S. Marine Mammal Commission, Washington, D.C., available from National Technical Information Service, Springfield, VA, PB-293 890, 343 pp. Geraci, J.R., Harwood, J., and Lounsbury, V.J., 1999, Marine mammal die-offs. Causes, investigations and issues, in Conservation and Management of Marine Mammals, Twiss, J.R., and Reeves, R.R. (Eds.), Smithsonian Institution Press, Washington, D.C., 367–395. St. Aubin, D.J., Geraci, J.R., and Lounsbury, V.J., 1996, Rescue, rehabilitation and release of marine mammals: An analysis of current views and practices, Proceedings of a workshop held in Des Plaines, Illinois, December 3–5, 1991, NOAA Technical Memorandum, NMFS-OPR-8, 65 pp. Seagers, D.J., Lecky, J.H., Slawson, J.J., and Sheridan Stone, H., 1986, Evaluation of the California Marine Mammal Stranding Network as a management tool based on record for 1983 and 1984, Administrative Report SWR-86-5, NMFS Southwest Region, Terminal Island, CA, 34 pp. Wilkinson, D.M., 1991, Report to Assistant Administrator for Fisheries: Program Review of the Marine Mammal Strandings Networks, U.S. Department of Commerce, NOAA, National Marine Fisheries Service, Silver Spring, MD, 171 pp. Wilkinson, D., and Worthy, G., 1999, Marine Mammal Stranding Networks, in Conservation and Management of Marine Mammals, Twiss, J.R., and Reeves, R.R. (Eds.) Smithsonian Institution Press, Washington, D.C., 396–411.



5 Marine Mammal Unusual Mortality Events Leslie A. Dierauf and Frances M. D. Gulland

Introduction The stranding of large numbers of marine mammals always commands a great deal of public, media, and scientific curiosity. Although these events occur with greater frequency along certain coastlines, they can occur worldwide, posing questions about their causes and potential effects on human health. Many animals stranding at one time is referred to as a mass stranding (see Chapter 6, Mass Strandings). When many animals strand over an extended period of time or in an unusual fashion, this is referred to as a Marine Mammal Unusual Mortality Event (MMUME). Providing humane care for the animals in such strandings, and determining the cause of such events are challenging tasks. Although identifying the immediate cause of such events is difficult, identifying predisposing factors and determining the effects of the event on the population dynamics and genetics of the remaining marine mammal population can be even more demanding (Harwood and Hall, 1990; Harwood, 1998; Baker, 1999). Causes of recent marine mammal die-offs and their investigations have recently been reviewed by Geraci et al. (1999). Although many investigations have been successful, each has its own set of complications and complexities and teaches different lessons (Geraci et al., 1999). As more reports are produced following investigations of MMUMEs, future responses will improve. To facilitate responses, and to maximize the chances for identifying the causes of unusual mortality events and their effects on marine mammal populations, a number of countries have developed contingency plans. In the United States, three specific events triggered the need for interested parties to develop a legal framework and subsequent law that addressed MMUMEs. The first was the Exxon Valdez oil spill in Prince William Sound, Alaska, in 1989 (Loughlin, 1994). The second was a stranding of 14 endangered humpback whales (Megaptera novaeangliae) off Cape Cod, Massachusetts in 1987 (Geraci et al., 1989), and the third was a bottlenose dolphin (Tursiops truncatus) die-off along the Atlantic seaboard between 1987 and 1988 (Geraci, 1989). In the 1st Session of the 102nd Congress, Congressman Walter Jones of North Carolina, who was Chairman of the Committee on Merchant Marine and Fisheries in the U.S. House of Representatives, introduced a bill called the “Marine Mammal Health and Stranding Act.” By late July 1992, the bill had passed out of committee, and a similar bill was moving through the Senate. On November 4, 1992, the Marine 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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Mammal Health and Stranding Response Act was signed into law by the President, and became Title IV of the Marine Mammal Protection Act (MMPA) (see Chapter 33, Legislation). In 1988, the dramatic phocine distemper epizootic that killed over 18,000 harbor seals (Phoca vitulina) in Europe raised awareness of the need for contingency plans to investigate marine mammal die-offs, and for long-term monitoring of strandings (Heide-Jorgensen et al., 1992; Thompson and Hall, 1993). In 1989, the Department of the Environment in the United Kingdom established a national program to investigate marine mammal mortalities in the United Kingdom and to coordinate responses. The sudden death of about 100 adult Hooker’s sea lions and over 1600 pups (Phocarctos hookeri) in the remote Auckland Islands off the southern tip of New Zealand in 1998 highlighted the need for preexisting sampling protocols and response plans. Although these have subsequently been developed, the lack of such plans at the time contributed to the difficulty in determining the predisposing factors that triggered the event (Baker, 1999). The Oxford English Dictionary defines the word contingency as a future event or circumstance where there is uncertainty of occurrence. Contingency plans are thus designed to guide responses during unusual events. These plans are imperative during MMUMEs, as such events are often sudden in onset, require early sampling to determine cause, are large scale, expensive to investigate, and command high public and media attention. This chapter reviews MMUMEs and the contingency plans in place to improve responses in the United States; Chapter 6 discusses mass strandings.

MMUME Responses in the United States To clarify protocols for response in the United States, strandings and MMUMEs have been clearly defined by law. A stranding (see Chapter 4, Stranding Networks; Chapter 6, Mass Strandings) is: • One or more marine mammals in the wild,

and • Dead on the beach or in the waters of the United States,

or • Alive and on the beach or shore, and —Either unable to return to the water, or —Although able to return to the water, is in need of medical attention, or —Unable to return to the water under its/their own power or without assistance.

Examples of stranding events are the regular and recurrent false killer whale (Pseudorca crassidens) mass strandings in Florida; the gray whale (Eschrichtius robustus) that becomes disoriented and caught up in a freshwater river; or the premature harbor seal (Phoca vitulina) pup that is abandoned by its mother. These are potential marine mammal mortalities, but they are not unusual. A MMUME is a stranding, but that stranding must: • Be unexpected; • Involve a significant die-off of any marine mammal population; and • Demand an immediate response.

Events deemed MMUMEs generally are caused by such things as geophysical catastrophic events, chemical spills, pollutant or contaminant discharges, biotoxins, microbial or parasitic


Marine Mammal Unusual Mortality Events

71

infections, and/or any other emergency affecting marine mammals in the wild. Recent examples of MMUMEs include the 1989 Exxon Valdez oil spill and sea otters (Enhydra lutris) in Alaska; the 1996 brevetoxicosis event in manatees (Trichechus manatus) off the west coast of Florida; and the 1998 domoic acid event in California sea lions (Zalophus californianus) along the California coast (Table 1).

The U.S. National Contingency Plan The United States has developed a contingency plan to respond to MMUMEs as mandated by Title IV of the Marine Mammal Protection Act. The purposes of Title IV are the following: 1. To bring together individuals with “knowledge and experience in marine science, marine mammal science, marine mammal veterinary and husbandry practices, and marine conservation, including stranding network participants”; 2. To establish a marine mammal health and stranding program and to set up a process within that program to facilitate the collection and dissemination of marine mammal health and health trend data, on marine mammal populations in the wild; 3. To help gather, collate, and correlate data on marine mammal health and marine mammal populations with data on physical, chemical, and biological environmental parameters, such as water sampling data from the Environmental Protection Agency (EPA), microbiological testing from the National Centers for Disease Control (CDC), weather data from the National Oceanic and Atmospheric Administration (NOAA), degree of habitat degradation, human disturbance, or food availability from the U.S. Fish and Wildlife Service (FWS); and 4. To provide coordinated and effective responses to unusual mortality events by establishing a mandated and timely process in which to act (MMPA, Title IV).

In addition, the processes within Title IV are designed to provide stranding network participants and marine mammal medical and conservation scientists with easily available broadbased data and reference materials. These reference materials are meant to be sufficient to help them better understand the connections between marine mammal health and the habitats upon which they depend for survival, as well as serve as general overall indicators of the health of our coastal and marine environs. The purpose of the MMUME National Contingency Plan is to outline actions that should be taken to protect public health and welfare; investigate and identify the cause of a mortality event, to minimize or mitigate the effects of a mortality event on the affected population, to provide for the rehabilitation of individual animals, and to determine the impact of a mortality event on the affected population. The FWS also has written an Oil Spill Response Contingency Plan (for wildlife in general), which is distributed through its Contaminants Program (USFWS, 1995).

Expert Working Group on MMUMEs Title IV established a decision-making body of scientific experts, called the Working Group on Marine Mammal Unusual Mortality Events (WGMMUME). The WGMMUME operates year round and meets once a year to coordinate efforts and apprise members of ongoing or past events. The group is composed of 12 experts from the fields of marine science, marine mammal science, marine mammal veterinary and husbandry practices, and marine conservation, including stranding network participants. A staff person from the National Marine Fisheries Service (NMFS) serves as executive director of the working group, and every 2 years, the working group chooses a chair from among its 12 members. Additional staff from the NMFS, the Marine Mammal Commission (MMC), and the FWS, and past members of the WGMMUME are welcome to

a

Common dolphins (Delphinus delphis)

1995

California sea lions (Zalophus californianus)

Mediterranean monk seals (Monachus monachus)

100

>150

28

Bottlenose dolphins (Tursiops truncatus) c

6

Right whales (Eubalaena glacialis)

b

∼150

Manatees (Trichechus manatus)

10

>200

220

2528

59

No. of Animals

FL panhandle, then MS, then AL, then LA Mauritania in Africa (western Sahara, southwest of Spain) North-central CA coast

Western North Atlantic

SW Coast of FL

Gulf of California (Sea of Cortez) Mexico Monterey Harbor, CA

Gulf Coast, TX

Coast of CA

Gulf Coast, TX

Location

Dx: Saxitoxin from dinoflagellate, Alexandrium, and/ or morbillivirus Dx: Leptospirosis

Unk; possibly red tide intoxication

Dx: Brevetoxin from the dinoflagellate (Gymnodinium breve) TDx: Ship strike and U.S. Navy underwater explosions

Unk

TDx: 18/25 dead dolphins exhibited morbillivirus TDx: Cyanide poisoning

Dx: Morbillivirus epizootic TDx: El Niño

Diagnosis

WG+, NOSC, leptospirosis occurs in California sea lions about every 4 years

Emaciated pups and juveniles, WG+, NOSC, NCP NOSC, NCP: in average year, fewer than 80 bottlenose dolphins strand here WG+; dead seabirds, too; cyanide found in dolphin liver and lung samples; source never identified WG+, NOSC, necropsies and testing for environmental contaminants negative WG+, OSC, IDST, R, toxic algal bloom; death via inhalation and ingestion report filed; 12% of 2/96 total manatee count WG+, NOSC, December to March, during winter calving season; 3 calves and 3 adults; skull fractures; abrupt deaths; eardrum ruptures WG+, NOSC, generally three or fewer strand in each of these areas; red tides and oyster bed closures WG; prior to event, total population only ~500 animals

WG+, OSC, NCP

Notes

Gulland et al., 1996

Osterhaus et al., 1997; Harwood, 1998; Hernandez et al., 1998

Bossart et al., 1998

Lipscomb et al., 1996

Lipscomb et al., 1996; Colbert et al., 1999

Reference

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1997

1996

Sea otters (Enhydra lutris)

Bottlenose dolphins (Tursiops truncatus) California sea lions (Zalophus californianus) Bottlenose dolphins (Tursiops truncatus)

1992

1994

Species

Year

TABLE 1 Marine Mammal Unusual Mortality Events since 1992



West coast of North America (Bering Sea to Baja Mexico) FL panhandle, in and near St. Joseph and St. Andrews Bays

Mid-Atlantic Coast (MA to NC)

Point Reyes, 20 miles north of San Francisco, CA Central CA coast

Dx: Brevetoxin from dinoflagellate (Gymnodinium breve)

Dx: Numerous causes, including decreased food availability, fisheries interactions, entanglement Unk

Unk; 3 of 85 were confirmed with sarcocystis meningitis Dx: Domoic acid intoxication from diatom (Pseudonitzschia australis)

WG+, NOSC, NCP, emaciation suggestive of nutritional disorder; variable chlorinated hydrocarbon levels WG−, large numbers of dead fish, birds, and sea turtles, as well

WG+, OSC, NCP, IDST, R, diatom cell counts reached 200,000/l; ingestion of sardines/anchovies; neurological signs, including seizures WG−, NCP, emaciated subadults

WG+, NOSC

Gulland, 2000; Scholin et al., 2000

Source: Table constructed from Marine Mammal Commission reports, 1992–1999.

Key: = contingency plan; CP NCP = no contingency plan; WG+ = WGMMUME decides it is a UME, requiring a response; WG− = WGMMUME decides it is not a UME, is within the normal range IDST = interdisciplinary scientific team participated in UME diagnostics; of variation for this particular species; R = scientific report written and filed/published in the scientific literature; WG+ = not a U.S. event; Dx = diagnosis made; OSC = on-site coordinator designated; TDx = tentative diagnosis only; NOSC = no on-site coordinator designated; Unk = cause unknown. a For mass die-offs prior to 1992, see Twiss and Reeves (1999, p. 376). b The northern right whale is the most endangered marine mammal in U.S. waters, and the most endangered large whale in the world, with only about 300 animals left in the population. c The Mediterranean monk seal is highly endangered.

87

Bottlenose dolphins (Tursiops truncatus)

216 (11 of them were alive; 55 carcasses were fresh)

273

Harbor porpoises (Phocoena phocoena)

1999

70

85

Gray whales (Eschrichtius robustus)

California sea lions (Zalophus californianus)

1998

Pacific harbor seals (Phoca vitulina)


Marine Mammal Unusual Mortality Events 73


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attend the annual meetings. Member terms are 3 years, with no person being allowed to serve more than two terms. Every 3 years, a third of the members rotates off, and new members are selected. The charges of the WGMMUME as mandated in Title IV are the following: • To determine whether or not a MMUME is occurring, • To determine after a MMUME has begun, when response to that MMUME is no longer necessary, and • To help develop a contingency plan for responding to MMUMEs.

The MMUME Response Details of the response to MMUMEs are given in the National Contingency Plan for Response to Marine Mammal Unusual Mortality Events (Wilkinson, 1996). To respond to a MMUME efficiently and effectively, there are several crucial elements that must be in place and operating: 1. A functional stranding network, with primary responders observing stranded marine mammals and reporting them to their regional stranding coordinator. The responders must provide precise information on the geographic location and approximate number and species of marine mammals involved. Each animal reported should have Level A data collected (Chapter 4, Stranding Networks; Chapter 21, Necropsy). 2. A regional coordinator, a national coordinator (from either the NMFS or the FWS, depending on the primary species involved in the UME), and a working group on MMUMEs, all of which work together according to the established plan. 3. A blueprint, plan, and protocols for animal rescue, rehabilitation and release, euthanasia, sample collection, referral laboratories to analyze collected samples, and long-term habitat and species protection. 4. Commitment and funding from the federal government to initiate a rapid response and to conduct complete investigations.

The response to a MMUME is shown in Figure 1. Each step of this process is essential for an effective response to proceed. Rapid and accurate information from each member of the stranding network to the regional stranding coordinator is the trigger for the process to begin. There are then two critical time constraints built into the MMUME response. First, the MMUME national coordinator is required to contact as many members of the working group as possible within 24 hours of a regional stranding coordinator contacting the NMFS. Second, members of the working group must call the MMUME national coordinator back immediately. Title IV does allow some flexibility if, at the request of any working group member, the MMUME coordinator needs to gather additional information on numbers, species, sexes, ages, and/or specific conditions associated with the MMUME to aid in decision making. Theoretically, the law states that each person in the working group within a maximum of 24 hours of obtaining the data needed must decide independently whether or not a MMUME is occurring and must register that decision with the MMUME coordinator. Once a majority of the working group has registered a yes or no vote, the MMUME coordinator announces whether (majority voted yes) or not (majority voted no) a MMUME is taking place. There are seven questions each expert working group member must ask: 1. Compared to historical records, is there a marked increase in the number of strandings of this species? 2. Are these marine mammals stranding at a time of year when historically strandings are unusual? 3. Are the increased strandings occurring in a localized area or over a wide geographic range, or is the event spreading geographically over time? 4. Is the species, age, or sex composition in the stranded animals different from what occurs normally in that geographic area or at that time of year?


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5. Are stranded animals exhibiting similar and/or unusual pathological changes or changes in general body condition from what is seen normally? 6. Are there animals alive in the area(s) where mortalities are occurring, and, if so, are they exhibiting any aberrant behaviors? 7. Does the stranding involve a critically endangered species?

Then, by law, unless time is needed to gather additional information as requested by any member of the working group, determination of whether or not an MMUME is occurring must

timeline Has the Regional Stranding Coordinator called the NMFS National MMUME Coordinator? 0 hours YES

NO

Process Stops

Has the NMFS National MMUME Coordinator called all the Members of the Working Group? 24 hours YES

NO

Contact NMFS Again

Has the NMFS National MMUME Coordinator received calls back from Working Group Members to be able to make a decision whether a MMUME is occurring?

YES

NO

Contact Working Group Again

Is a MMUME occurring?

YES 48 hours

NO

Process Stops Regional Stranding Coordinator, Continues to Watch, and Keeps Regular Contact with NMFS MMUME Coordinator

MMUME National Coordinator informs Regional Stranding Coordinator a MMUME is occurring MMUME National Coordinator through Secretary of Commerce designates On-Site Coordinator MMUME National Coordinator transfers responsibility for action to the On-Site Coordinator On-Site Coordinator makes immediate recommendations to the Regional Stranding Coordinator on how best to proceed with response activities On-Site Coordinator takes over response, following the Contingency Plan to the best of his/her abilities, utilizing professional judgment, and assembles response team and plan On-Site Coordinator or his/her designee remains on site at MMUME coordinating the response FIGURE 1 Flowchart and timing of response to MMUME in the United States.


76


On-Site Coordinator

Live/Dead Animal Rescue Response

Legal Counsel and National MMUME Coordinator

Live/Dead Animal Research Response Activities

Command Operations and Administrative Response

FIGURE 2 Coordinated team response interactions during a MMUME. (Adapted from the U.S. National Contingency Plan.)

take place within 48 hours of a regional stranding coordinator contacting the NMFS about a possible event. If the working group believes a MMUME is indeed occurring, an appropriately qualified onsite coordinator (OSC) is immediately designated to mobilize and manage the national response to the event. Depending on the species involved and the location of the MMUME, the OSC will be either a NMFS or a FWS regional director or an individual designated by that regional director. Because the OSC is responsible for directing the response, the individual must have strong management and leadership capabilities, highly effective communication skills, the capacity to make decisions with minimal use of intermediaries, the ability to access information and expertise including interagency expertise, and a familiarity with the contingency plan and the stranding network. The OSC is also responsible for preparing a report containing results of scientific investigations and recommendations for subsequent monitoring and/or management activities. The coordination of team efforts once an on-site coordinator has been designated for a MMUME is shown in Figure 2. Through the National Contingency Plan, adequate funding, personnel for the team, and logistical support, such as ships, aircraft, and other heavy equipment, are made available to carry out an efficient and effective response, whether the marine mammal involved in the MMUME is under NMFS or FWS jurisdiction (see Chapter 33, Legislation).

MMUME Fund Title IV established an interest-bearing account in the Federal Treasury called the “Marine Mammal Unusual Mortality Event Fund” to be used exclusively for costs associated with preparing for and responding to MMUMEs, which remains available until expended. Monies provided to the fund come from multiple sources, including Congressional appropriations, special funds appropriated to the Secretary of Commerce, and monies received by the U.S. government in the form of public or private gifts, devises, and/or bequests. The acceptance and solicitation of donations into a fund such as this is highly unusual in the federal government, but allowable and anticipated under Title IV of the MMPA.



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Anyone wishing to donate funds to the MMUME Fund is asked to contact the NMFS or the chair of the working group. Donations can be sent directly to NMFS, 1335 East-West Highway, Silver Spring, MD 20910, with a notation attached that the money is to be used “exclusively for marine mammal MMUME through the MMUME Fund.” If every person reading this chapter sent just $5 each, the fund would grow incrementally and be able to support the important tasks and responses needed to continue to make the MMUME program successful. Although the fund is coordinated by the NMFS, it is available for response to any MMUME, including those under NMFS and FWS jurisdiction.

Lessons Learned The Cooperative Response Stranding network participants are highly vigilant in alerting federal officials whenever there is even an inkling of a MMUME. Scientists and stranding network participants give maximum effort in reacting to MMUMEs and in providing tissues and samples for furthering knowledge of MMUMEs in general and of individual MMUMEs in particular. Facilities must, as new volunteers arrive to assist, make all stranding network volunteers aware of national plans and needs. Participants must understand their reporting obligations and the importance of Level A data (see Chapter 4, Stranding Networks; Chapter 21, Necropsy). All original members of the working group have now been replaced through attrition, and the working group under the directorship of its chair continues to be highly productive, developing standardized protocols, assisting with developing new contingency plans and revising existing plans, and devising strategies to increase funding for MMUME responses. Plans are being developed for MMUMEs that recur, such as leptospirosis, El Niño events, and domoic acid toxicity in California sea lions off the West Coast of the United States. Interdisciplinary scientific and logistical teamwork is important to obtain diagnoses. In the last few years, each MMUME in the United States and elsewhere has garnered a response from a multitude of players in the scientific community, a kind of collaborative response rarely seen in the past. Federal, state, regional, stranding network, and private agencies and individuals participate, as do many academic institutions. The scientific and gray literature associated with MMUMEs now is written by multiple scientific contributors. Interagency cooperation has improved. The NMFS, the U.S. Geological Survey, the EPA, and the FWS met in October 1998 and decided to create an interagency working group to address the uncertainties and unknowns regarding contaminant levels that are being detected in marine mammals. Although the NMFS, the FWS, and the EPA do not yet work seamlessly together, there has been noticeable improvement since Title IV of the MMPA came into existence. Around the world, national contingency plans to respond to unusual mortality events in marine mammals are under development or under discussion. The UN Environment Program (UNEP) has an action plan for marine mammals worldwide. Although lack of funding at any particular time can hinder the magnitude of a response anywhere at any time, it is the unending support of the volunteers in stranding networks worldwide that makes the response possible and successful. The WGMMUME has assisted the NMFS and the FWS in developing and releasing a series of contingencies plans, including the National Contingency Plan for Response to Marine Mammal Unusual Mortality Events (Wilkinson, 1996), and the Contingency Plan for Catastrophic Rescue and Mortality Events for the Florida Manatee and Marine Mammals (Geraci and Lounsbury, 1997). In addition, the NMFS is working on a new contingency plan for the Hawaiian monk seal.


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The Process In the United States, not all stranding network members or participants are aware of the MMUME law or process, or of the existence of a national contingency plan. Communication between the federal agencies and the working group must be as rapid as possible, as does the response of the working group. Members of the working group must make their individual decisions about whether or not a MMUME is occurring within 48 hours, so the response time is effective. Single, local response teams at the stranding network level cannot be left to respond on their own to huge, time-consuming MMUMEs without the aid of personnel or funding from the federal government.

UMMME Fund The NMFS and the FWS always are concerned about funding constraints in trying to implement their programs relating to marine mammals. Funding is important because it supports the following efforts: • Communication, helping staff, who at times can feel overburdened with excessive workloads; • Baseline data collection and collation, including information on stranding rates, disease, and environmental contaminants for use in securing diagnoses of MMUME causes; • MMUME sample/tissue data collection, archiving, and analysis; and • Rapidity of the response to MMUME.

A 1994 Congressional amendment to the MMPA allows monies from the MMUME Fund to be used for care and maintenance of marine mammals seized by NMFS or FWS agents when the level of care the animals are receiving is inadequate. This seizure is important to marine mammal well-being, but is not a MMUME, and original Congressional intent was never to use the fund for such purposes. The intent was always to use the fund for wild marine mammals and not for animals held in captivity at aquaria, zoos, or other U.S. facilities (U.S. House of Representatives, 1992). Thus, it is extremely important when making donations to the MMUME Fund that the NMFS be instructed that the money is to be used “exclusively for marine mammal UME.”

Results Accrued from Title IV of the MMPA There has been definite improvement in the collection quantity and quality of marine mammal disease data. More final diagnoses have been made since passage of Title IV, although the predisposing factors often remain unclear. It is the authors’ hope that in the future there will be more integration of baseline health, population parameters, and ecosystem changes with investigations of MMUMEs. This will help determine whether or not there are real long-term alterations occurring in ocean health, as suggested by Harvell and co-workers (1999), rather than simply improvements in detection and reporting. Relative to a response to unusual stranding events prior to 1992, there is now a coordinated effort, with much interaction among federal, state, regional, and local participants. Funding for MMUME responses and tissue analyses, as well as database establishment and maintenance, is critical. The more people who know about MMUMEs and Title IV, and the more people who have a passion for marine mammal and ecosystem health, the more people there will be to lobby Congress and their individual Senators and Representatives to ensure that annual appropriations are provided for the program. Private donations and gifts are welcome also.



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How Can You Help? Volunteer with local stranding networks on a regular basis. Understand the plans and legislation in place to facilitate responses to dead marine mammals (see Chapter 4, Stranding Networks; Chapter 6, Mass Strandings; Chapter 33, Legislation). Donate supplies and funds to support local efforts. Help tackle logistical problems facing stranding network participants during investigations. Assist with administrative and communication tasks, as well as with the more attractive jobs working directly with the animals. Send gifts and donations to the national fund for MMUMEs. Tell everyone you know about MMUMEs and how we can learn more from responding quickly to them and working together to determine and explain the causes of MMUMEs. In your research endeavors, keep marine mammal health and well-being in the forefront, developing rapid, sensitive, and specific tests for diagnosing disease and finding new and effective ways to treat marine mammals found alive during MMUMEs. Always consider factors beyond conventional clinical medicine when dealing with wild animals—environmental changes, population dynamics, and genetics.

Conclusion Unusual mortality events and other marine mammal strandings are effective learning tools for diagnosing factors affecting the health of marine mammal populations. If a marine mammal is still alive or freshly dead, tissues can be collected, using a standardized set of methodologies for quality-controlled analysis. The results may lead to an explanation of what caused the individual or group of marine mammals to strand. Even more importantly, placing these data in a national, accessible database will allow information from one event to be compared with that from another. All of this information can be compared with reference materials taken from nonstranding marine mammals in the wild. Such carefully planned procedures will provide the most insightful evidence for determining why marine mammals strand, how MMUMEs occur, and when these events are harmful to marine mammal populations and the ecosystems upon which they depend. Marine ecosystems worldwide are being negatively impacted by multiple factors, and they need immediate attention. Only by concentrating everyone’s attention on marine mammals and the habitats in which they live, will we be able to continue to be fascinated and mesmerized by healthy marine mammals in the wild for generations to come.

Acknowledgments The authors thank Mona Haebler and Tom O’Shea for their reviews of this chapter. Both have served on the WGMMUME, as have the authors.

References Baker, A., 1999, Unusual mortality of the New Zealand sea lion Phocarctos hookeri, Auckland Islands, January–February 1998, Report of a workshop held 8–9 June 1998, Wellington, NZ, and a contingency plan for future events, New Zealand Department of Conservation, 84 pp. Bossart, G.D., Baden, D.G., Ewing, R.Y., Roberts, B., and Wright, S., 1998, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizootic: Gross, histologic and immunohistologic features, Toxicol. Pathol., 26: 276–282. Colbert, A.A., Scott, G.I., Fulton, M.H., Wirth, E.F., Daugomah, J.W., Key, P.B., Strozier, E.D., and Galloway, S.B., 1999, Investigation of unusual mortalities of bottlenose dolphins along the midTexas coastal bay ecosystem during 1992, NOAA Technical Report NMFS 147, U.S. Department of Commerce, Seattle, Washington, 23 pp.


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Costas, E., and Lopez-Rodas, V., 1998, Paralytic phycotoxins in monk seal mass mortality, Vet. Rec., 142: 643–644. Geraci, J. R., 1989, Clinical investigation of the 1987–1988 mass mortality of bottlenose dolphins along the U.S. central and south Atlantic coast, Final Report, U.S. Marine Mammal Commission, Washington, D.C., 63 pp. Geraci, J.R., and Lounsbury, V.J., 1997, Contingency plan for catastrophic manatee rescue and mortality events, Florida Department of Environmental Protection, Florida Marine Research Institute, Contract Report MR 199, 136 pp. Geraci, J.R., Anderson, D.M., Timperi, R.J., St. Aubin, D.J., Early, G.A., Prescott, J.H., and Mayo, C.A., 1989, Humpback whales (Megaptera novaeangliae) fatally poisoned by dinoflagellate toxin, Can. J. Fish. Aquat. Sci., 46: 1895–1898. Geraci, J.R., Harwood, J., and Lounsbury, V.J., 1999, Marine mammal die-offs. Causes, investigations and issues, in Conservation and Management of Marine Mammals, Twiss, J.R., and Reeves, R.R. (Eds.), Smithsonian Institution Press, Washington, D.C., 367–396. Gulland, F., 2000, Domoic acid toxicity in California sea lions (Zalophus californianus) stranded along the central California coast, May–October 1998, NOAA Technical Memorandum, NMFS-OPR, 17, 45 pp. Gulland, F.M.D., Koski, M., Lowenstine, L.J., Colagrass, A., Morgan, L., and Spraker, T., 1996, Leptospirosis in California sea lions (Zalophus californianus) stranded along the central California coast, 1981–1994, J. Wildl. Dis., 32: 572–580. Harvell, C.D., Kim, K., Burkholder, J., Colwell, R.R., Epstein, P.R., Grimes, J., Hofmann, E.E., Lipp, E.K., Osterhaus, A.D.M.E., Overstreet, R., Porter, J.W., Smith, G.W., and Vasta, G.R., 1999, Emerging marine diseases—climate links and anthropogenic factors, Science, 285: 1505–1510. Harwood, J., 1998, What killed the monk seals? Nature, 393: 17–18. Harwood, J., and Hall, A., 1990, Mass mortality in marine mammals: Its implications for population dynamics and genetics, Trends Ecol. Evol., 5: 254–257. Heide-Jorgensen, M.P., Harkonen, T., Dietz, R., and Thompson, P.M., 1992, Retrospective of the 1988 European seal epizootic, Dis. Aquat. Organisms, 13: 37–62. Hernandez, M., Robinson, I., Aguilar, A., Gonzalez, L.M., Lopez-Jurado, L.F., Reyero, M.I., Cacho, E., Franco, J., Lopez-Rodas, V., and Costas, E., 1998, Did algal toxins cause monk seal mortality? Nature, 393: 28–29. Lipscomb, T.P., Kennedy, S., Moffett, D., Krafft, A., Klaunberg, B.A., Lichy, J.H., Regan, G.T., Worthy, G.A.J., and Taubenberger, J.K., 1996, Morbilliviral epizootic in bottlenose dolphins of the Gulf of Mexico, J. Vet. Diagn. Invest., 8, 283–290. Lipscomb, T.P., Schulman, Y.D. Moffett, D., and Kennedy, S., 1994, Morbilliviral disease in Atlantic bottlenose dolphins (Tursiops truncatus) from 1987–1988 epizootic, J. Wildl. Dis., 30: 567–571. Loughlin, T.R. (Ed.), 1994, Marine Mammals and the Exxon Valdez, Academic Press, San Diego, CA, 395 pp. MMC, Marine Mammal Commission, 1992–1999, Annual Reports to Congress, Bethesda, MD, available January of each following year. MMPA, Title IV, Marine Mammal Protection Act of 1972, as amended, 1995, 16 USC 1421 ff. Osterhaus, A., Groen, J., Neisters, H., Van de Bildt, M., Vedder, B.M.L., Vos, J., van Egmond, H., Sidi, B.A., and Barham, M.E.O., 1997, Morbillivirus in monk seal mass mortality, Nature, 388: 838–839. Scholin, C.A., Gulland, F., Doucette, G.J., Benson, S., Busman, M., Chavez, F.P., Cordaro, J., DeLong, R., De Vogelaere, A., Harvey, J., Haulena, M., Lefebvre, K., Lipscomb, T., Loscutoff, S., Lowenstine, L.J., Marin III, R., Miller, P.E., McLellan, W.A., Moeller, P.D.R., Powell, C.L., Rowles, T., Silvagni, P., Silver, M., Spraker, T., Trainer, V., and Van Dolah, F.M., 2000, Mortality of sea lions along the central California coast linked to a toxic diatom bloom, Nature, 403: 80–84. Thompson, P.M., and Hall, A.J., 1993, Seals and epizootics—what factors might affect the severity of mass mortalities? Mammal Rev., 23: 149–154. USFWS, U.S. Fish and Wildlife Service, 1995, Oil Spill Contingency Plan, 1995.



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U.S. House of Representatives, Marine Mammal Health and Stranding Response Act, Committee Report, 1992, Report 102-758, July 30, 14 pp. Wilkinson, D.M., 1996, National Contingency Plan for Response to Unusual Marine Mammal Mortality Events, NOAA Technical Memorandum NMFS-OPR-9, 9/96, Silver Spring, MD.


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6 Mass Strandings of Cetaceans Michael T. Walsh, Ruth Y. Ewing, Daniel K. Odell, and Gregory D. Bossart

Introduction A mass stranding of cetaceans is an event in which two or more individuals of the same species, excluding a single cow–calf pair, beach within a given spatial and temporal reference (Wilkinson, 1991). A mass stranding event may span 1 or more days and range over miles of shoreline, bridging multiple counties, or sandbars and outlying keys. A variety of species have been affected; Odell (1987) listed 19 odontocete species known to mass-strand. Aristotle recorded sightings of stranded cetaceans 2300 years ago. Cetaceans continue to mass-strand, yet the causes of the majority of these events remain unclear. Mass strandings have received more attention as coastal human populations increase, making discovery of stranded animals more likely. Documentation of stranding events has improved over the last 70 years, the earliest organized attempts originating in England. These records have allowed reviews of such occurrences (Fraser, 1934; 1946; 1953; 1956; Geraci, 1978; Sergeant, 1982). Despite the attention mass strandings receive from the public and scientific community alike, they remain hard to manage, and the reasons for their occurrence remain hard to identify. Geraci et al. (1999) produced an excellent review of marine mammal die-offs, summarizing various etiologies of mass-stranding events. Table 1 lists a compilation of mass strandings, mostly from the Smithsonian marine mammal database and the Southeast United States (SEUS) marine mammal stranding network database, that have occurred along the East Coast of the United States within the past 12 years (1987 through 1999). Causes of most of these events are either unknown or ambiguous, theories being supported only by circumstantial evidence.

Theories to Explain Mass Strandings As long as people have been aware of mass strandings, theories have been formulated to explain why marine mammals mass-strand on beaches (Dudok Van Heel, 1962; Geraci et al., 1976; Eaton, 1979; 1987; Geraci and St. Aubin, 1979; Odell et al., 1980; Best, 1982; Cordes, 1982; Wareke, 1983). Anecdotal theories for why whales strand include that these species whose ancestors were land mammals have an evolutionary memory compelling them back to land, that the animals are distressed and/or in pain and are committing suicide, and that they are avoiding drowning. Other more accredited theories include that sloping beaches give poor sonar reflection 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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TABLE 1 Mass Strandings along the East Coast of the United States from 1987 through 1999 Species

Year

Month

Day

Number of Animals

State

Ref.

P. crassidens D. delphis L. acutus K. breviceps L. acutus L. acutus S. bredanensis G. macrorhynchus T. truncatus D. delphis L. acutus L. acutus F. attenuata S. coeruleoalba P. crassidens K. breviceps L. acutus P. macrocephalus L. acutus G. melas G. griseus S. coeruleoalba G. macrorhynchus G. macrorhynchus S. bredanensis G. macrorhynchus G. melas G. melas G. melas G. melas G. melas G. macrorhynchus G. macrorhynchus G. macrorhynchus F. attenuata Z. cavirostris (?) F. attenuata L. acutus K. breviceps F. attenuata S. clymene G. melas T. truncatus D. delphis S. frontalis L. acutus S. attenuata G. macrorhynchus G. griseus K. breviceps D. delphis

1987 1987 1987 1987 1987 1987 1987 1987 1987 1988 1988 1988 1988 1989 1989 1989 1989 1990 1990 1990 1991 1991 1991 1991 1991 1991 1991 1991 1991 1991 1991 1992 1992 1992 1992 1992 1992 1992 1992 1992 1992 1992 1992 1993 1993 1993 1993 1993 1993 1993 1993

1 2 3 8 9 9 10 11 12 2 4 4 5 1 7 8 8 4 8 12 1 3 3 4 4 7 9 9 9 10 12 1 2 2 3 6 7 8 8 9 12 12 12 1 3 4 9 11 11 11 12

2 4 7 23 5 5 18 14 1 4 29 30 7 26 11 9 30 19 9 11 20 9 b 24 –30 b 11–20 24 b 21–22 9 10 29 8 24 30 10 15 30 25 3 27 31 4 6–10 12 13 1 15 6 6 3 20 21 20

6 5 3 3 20 10 3 29 3 5 3 3 4 3 3 3 4 5 9 53 3 b 4 /5 27 12 10 11 32 27 17 16 31 13 3 8 2 3 2 6 3 3 23 19 6 6 2 8 5 6 5 2 4

LA MA MA FL ME MA FL FL SC MA MA MA GA MA FL NC ME FL ME MA NC FL FL FL FL FL MA MA MA MA MA FL FL FL FL FL FL MA FL FL FL MA MA MA MS MA FL FL MA FL MA

a a a a; b a a a; b a; b a a a a a; b a a; b a a a; b a a; c a; b a; b a; b a; b a; b a; b b; c c a; c a; c a; c a; b a; b a; b b a; b b a; c b b b a; c a a b c a; b b a a c


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Mass Strandings of Cetaceans

TABLE 1 Mass Strandings along the East Coast of the United States from 1987 through 1999 (continued) Species G. macrorhynchus G. macrorhynchus D. delphis L. acutus L. hosei L. acutus K. breviceps L. acutus L. acutus G. macrorhynchus S. clymene G. macrorhynchus G. macrorhynchus G. macrorhynchus G. macrorhynchus F. attenuata K. breviceps S. attenuata G. macrorhynchus L. acutus L. acutus S. bredanensis D. delphis G. macrorhynchus G. macrorhynchus L. acutus D. delphis S. bredanensis M. europaeus G. melas S. bredanensis L. acutus G. macrorhynchus S. attenuata L. acutus S. bredanensis Z. cavirostris

Year 1994 1994 1994 1994 1994 1994 1994 1994 1995 1995 1995 1995 1995 1995 1995 1995 1995 1996 1996 1997 1997 1997 1997 1998 1998 1998 1998 1998 1998 1998 1998 1999 1999 1999 1999 1999 1999

Month 2 b 2–3 3 3 7 10 11 12 1 3 6 7 8 8 9 9 12 1 5 5 8 12 11 1 1 1 1 2 8 11 12 3 5 8 8 8 10

Number of Animals

Day b

17–24 b 26–24 5 14 13 9 5 30 4 24 15 1 15 21 15 16 11 16 31 28 12 14 16 3 12 c 29 /31 31 4 28–31 6 28 19 5 2 11 21 3

46 b 4/3 3 6 b 30/28 7 4 23 12 2 18 32 4 9 7 5 b 6/3 11 2 2 2 34 10 7 8 c 97/82 16 2 9 2 12 50 2 3 6 5 4

State

Ref.

FL NC MA MA FL MA NJ MA MA NC FL FL FL FL FL VI FL FL FL MA MA FL MA FL FL MA MA FL NC FL FL MA FL FL MA GA VI

a; b a; b a; c a; c a; b a a a; c c b a; b a; b b a; b a; b a a; b a; b b a a a; b a; c a; b a; b a; c c b b b a; b c b b c b b

Note: (?) indicates species uncertain in database record. Shaded individual species records have been considered to be from the same mass stranding event; however, they have been recorded as separate events within the referenced databases. a Refers to data within the Cetacean Distributional Database, Smithsonian Institute. b Refers to data in the SEUS marine mammal stranding network database. c Refers to data referenced in Wiley et al., in review.

which misleads the animals ashore; that geomagnetic disturbances affect their ability to navigate geomagnetically; that acoustic navigation is lost as a result of parasitic destruction of the eighth cranial nerve; that coastlines are unfamiliar to the animals; that the animals strand as a result of geologic disturbances, such as earthquakes or underwater volcanoes; and that mass strandings involve pelagic species, which may have difficulty navigating in shallow waters.


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It is likely that many species involved in mass strandings use geomagnetic cues to migrate (Kirschvink et al., 1985; Klinowska, 1985a,b). Klinowska (1985b) proposed geomagnetic disturbances as an explanation for live strandings in the United Kingdom. This theory is based on the study of coastline geomagnetic maps and the finding that correlations exist between stranding sites and relative intensity of the local geomagnetic fields. It is likely that this theory is a factor in explaining where animals strand, rather than why they strand, and it is certainly possible that a group of ill individuals will overlook other sensory modalities and ultimately follow geomagnetic or shoreline clues into a specific location. This may be a partial explanation for why certain beaches, such as on Cape Cod, Massachusetts, seem to experience repeated mass-stranding events. The loss of acoustic navigation ability (“sonar”) as a result of parasitic involvement may explain some mass strandings (Ridgway and Dailey, 1972). Parasites are common in wild species (see Chapter 18, Parasitic Diseases), and their presence in locations such as the middle or inner ear could lead to disorientation. Morimitsu et al. (1986) demonstrated eighth cranial nerve destruction induced by Nasitrema spp. at the junction with the inner ear in three cetacean species. However, there is some question about the validity of these conclusions, as it was stated in a subsequent publication that these specimens were not fresh, and freeze artifact may have affected the histological appearances of the tissues (Morimitsu et al., 1987). The lack of early evidence for specific viral or bacterial etiologies in some stranding events in the mid-1980s reawakened the discussion of the role of pod cohesion as a major factor in mass strandings. In 1986, during a mass stranding of false killer whales (Pseudorca crassidens) in the Florida Keys, the influence of social structure was plainly illustrated (Walsh et al., unpubl. data). After repeatedly stranding and being pushed back to sea by the public, a group of false killer whales eventually stranded in the Florida Keys (Odell et al., 1980). The group of 30 animals was spread over more than 12 miles along shallow waters and numerous islands. The effort to coordinate and relocate the surviving 16 animals to a central location resulted in the youngest and smallest animals being moved first to a small isolated bay. At first these five young animals were actively swimming and investigating the shallow bay. They appeared confused, but they were active. When one of the larger adult male animals was transported into the bay, he immediately beached himself on one edge of the shore. Each of the younger animals then lined up neatly beside him and did not move from his side. Whether the response was based on visual or auditory cues was unknown, but as each animal was added to the group, this response was repeated until all survivors were in one line.

Current Investigations into Mass Strandings Investigations of mass-stranding events have evolved and continue to evolve as more standardized approaches are applied. For example, a mass stranding of Atlantic white-sided dolphins (Lagenorhynchus acutus) yielded valuable information on pathological conditions that were present, including parasite identification and numbers, along with other baseline life history data (Geraci and St. Aubin, 1977). In a subsequent mass-stranding investigation in 1986 involving shortfinned pilot whales (Globicephala macrorhynchus), clinical pathology was emphasized. Blood samples for complete blood counts (CBC) and serum chemistries were taken from all live animals to elucidate observed clinical symptoms of disease (Walsh et al., 1991). The diagnostic workups also included cultures of the respiratory, reproductive, and gastrointestinal systems. Serum was initially used for serological analyses for certain known domestic animal and marine mammal pathogens; however, serum subsamples were also archived for future retrospective analyses. At necropsy, samples were collected for histopathological and toxicological analyses, urinalysis, and various additional tissue cultures (Bossart et al., 1991). This investigation, while comprehensive, was limited by three factors: interest/disciplinary focus, response crew abilities, and finances.


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Although there was evidence of illness in individuals from these mass strandings, no specific etiology for the stranding event was identified. New issues have been raised as each incident is more thoroughly investigated. What infectious agents, such as viruses or bacteria, may be involved? What role do anthropogenic or naturally occurring biotoxins or contaminants play in mass strandings? What factors are primary; which are secondary? Could the original problem, which may have occurred weeks or months before, out at sea, be missed?

Evaluation of a Mass Stranding One approach to evaluating a stranding event is given in Table 2. This approach includes assessments of environmental conditions and trends, the group of animals as a whole, and the individuals of that group. The environmental evaluation should list all potential factors, including: 1. Previous strandings at this site (historical perspective); 2. Geomagnetic maps (if available); 3. Topographic and bathymetric characteristics and anomalies (beach type, slope, presence of barrier islands, sandbars, landslides, volcanic eruptions, earthquakes); 4. Tide factors, sea surface temperature, salinity, fronts, currents, and other oceanographic factors; 5. Storms within the last few weeks; 6. Available local fishing data on local fishery changes; 7. Algal blooms; 8. Toxic material spills; 9. Acoustic events; and 10. Other species mortalities.

Evaluation of animal groups should include: 1. Recognition that in some species of cetaceans there are strong social ties between group members, which may result in individuals blindly congregating around ill leaders or other ill individuals; thus, the species involved, and the leader (if possible) should be identified; 2. Group demographics (sex and age distribution); 3. The ratio of live to dead animals; 4. Cow–calf pairs; and 5. Evaluation of individuals involved. TABLE 2 Factors to Evaluate during a Mass Stranding Environmental Local Adverse weather: Storms Beach Topography Previous stranding history Current and tides Acoustic events: Land slides Volcanic eruption Underwater experiments Anthropogenic noise

Cetacean Regional

Group

Individual

Weather pattern shift: El Niño La Niña Foodborne toxins Food availability Harmful algal blooms Oil spill Pesticide runoff

Social bonds: Leader illness Cow–calf pairs Breeding season: Pregnant females Infectious disease: Acute process Chronic disease

Appearance Attitude Heart rate and character Respiratory rate and character Hematology and serum biochemistry


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Management of a Mass Stranding Strandings are generally very complicated events. Proper management requires experienced, organized rescue efforts, including individuals trained in stabilizing live animals, rapidly diagnosing illnesses, and arranging for possible extended rehabilitation of ill animals. In some cases, controlling interference from untrained individuals is also a priority. To work in concert with local law authorities, such as the Marine Patrol or local police, members of the mass-stranding rescue team should make contact with the law enforcement officer in charge. A temporary plan (which may include aerial survey and observations) should be implemented to determine the number of animals involved, where they are located, the accessibility of the stranding location and to evaluate other pertinent circumstances (Figure 1). If the animals are spread over a large area, it may be advisable to consolidate the individual animals (weather permitting) into one location. If there is adequate help available, individuals are assigned to each animal to provide temporary first aid, including keeping the animal sternal to avoid inhalation of debris. Animals exposed to sunlight must be kept moist, cool, and shaded. Zinc oxide can be applied to briefly towel-dried skin, to help deflect sunlight and decrease sunburn. Pouring water over the animal’s body will also help keep the skin from drying and the animal from overheating. If towels are placed over the animal, they must be kept wet and not placed where they may occlude respiration. All individual animals should be identified with tape or tags (such as small spaghetti tags or roto tags) (see Chapter 38, Tagging and Tracking) placed in the dorsal fin to facilitate correlation between clinical and pathological data collection, as well as later identification should the animals be released and re-strand. Algorithms to aid in evaluations of individuals within the group are summarized in Figures 2 and 3. These flowchart approaches to individual evaluations involve on-site monitoring of

Verification

Site Evaluation

Accessible

Evaluate Group

Inaccessible

Return to Sea

See Figure 2

FIGURE 1 Algorithm for initial mass stranding response.

Euthanasia


89


Verified Stranding Accessible

All Alive

Evaluation and Triage

Alive and Dead

Alive

All Dead

Dead

Keep Sternal Heart Rate Respiratory Rate Physical Exam Blood Sample

See Figure 3

Necropsy

Field Data Level A Data Cetacean Data Other Data Measurements Photos

Necropsy Tissues Cultures

FIGURE 2 Algorithm for evaluation of animals that are accessible.

health status and separation of affected individuals into groups, based on clinical findings, which include (1) those likely to survive; (2) those apparently stable, but showing obvious signs of illness; and (3) those unlikely to survive. Individual health monitoring needs to include heart rate, respiratory rate, and attitude. Heart rates can be monitored in a partially submerged animal by placing the hand on the area between the pectoral flippers, and feeling for the reverberations of the heart through the chest wall. For safety reasons, this procedure should not be attempted with struggling or very large animals. In totally beached animals, which are lying laterally (although some animals beach sternally), heart rate may be visualized by movement of the sternal area. In a mass stranding of 30 false killer whales in Florida, heart rates ranged from 60 to 150 beats per minute (bpm) (Walsh et al., unpubl. data). Normal heart rates of this species are approximately 60 to 100 bpm and respiratory rates are 8 to 18 breaths per 5 min. The animals that lived the longest were five animals with near normal heart and respiratory rates (Walsh et al., unpubl. data). In addition to physical information, blood samples should be taken from each individual before any treatments are given. Blood collection is discussed elsewhere (see Chapter 19, Clinical


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Blood Analysis and Physical Exam Results

Normal

Abnormal

Release Rehabilitation

Rehabilitation

Survival (>6 mo)

Death

Euthanasia

Necropsy

Field data Release

Retain

Radio-tag or Mark

Tissues Cultures

FIGURE 3 Algorithm for animal evaluation and disposition.

Pathology). Care must be taken when sampling stranded cetaceans, because they are capable of inflicting injury with their flukes, especially to inexperienced volunteers. At a minimum, blood sample volume should be sufficient to include CBC, serum chemistries, and serum electrolyte levels; however, optimally, additional serum is required for additional diagnostic analyses and for archival purposes. It is often possible to have pertinent tests run on an emergency basis utilizing local hospitals and veterinary clinics close to stranding sites. Emergency clinical laboratory tests should include manual packed cell volume, refractometer-determined total protein, fibrinogen, white blood cell count, glucose, blood urea nitrogen, creatinine, calcium, and electrolytes. These tests can aid the on-site clinician and rescue crew making decisions regarding the disposition of the group. Any residual serum and EDTA plasma should be retrieved from the hospitals and/or veterinary clinics and archived for future analyses. Fibrinogen tests require special tubes containing sodium citrate, and need to be spun, plasmaseparated, and analyzed or frozen in plastic vials within 1 hour of sampling to ensure accuracy. If possible, a centrifuge should be available on site to allow serum or plasma separation as soon as possible. New handheld, portable analyzers are available to analyzed some electrolytes, chemistries, and blood gas parameters on site. Blood glucose monitors may also be helpful in evaluating animals. Biochemical and hematological abnormalities found in individuals of each stranding may vary widely. In the stranded false killer whales, pod abnormalities included hemoconcentration, leukopenia, elevated liver enzymes, hypernatremia, hyperchloremia, and hypocalcemia (Table 3).

0.2 0.2 0.3 0.1 0.3 0.2 0.4 — 1.0 0.3 0.3 0.4 0.3 0.2

111 88 96 135 122 115 131 — 128 122 154 99 138 280

26 92 74 113 25 166 53 — 74 25 58 139 — 65

6.4 6.3 5.5 5.3 5.7 6.4 7.5 — 6.9 5.7 5.9 4.5 6.5 7.2

TP g/dl 2.7 2.3 3.1 2.9 2.8 2.8 3.1 — 3.2 2.8 3.0 2.0 3.0 3.6

Alb g/dl 3.7 4.0 2.4 2.4 2.9 3.6 4.4 — 3.7 2.9 2.9 2.5 3.5 2.8

Glob g/dl 9 14 32 20 49 22 12 47 8 49 17 30 11 14

Amy U/l 239 166 — 240 440 317 — 317 250 208 76 — — —

Lip U/l 106 106 363 269 159 66 108 56 158 159 201 479 160 242

AP U/l 112 59 3 15 80 40 9 60 105 80 30 38 33 15

ALT U/l 675 1490 423 279 1080 655 490 603 >2500 1080 382 740 830 110

AST U/l 20 21 — 27 27 29 — 30 16 28 19 — — 26

GGT U/l

787 281 155 104 498 984 677 331 1174 498 606 1205 535 60

CK U/l

2692 1089 567 380 1258 980 1517 1083 725 1258 725 1054 1546 382

LDH U/l

9.0 6.8 7.0 6.8 6.6 7.6 7.6 — 8.3 6.6 7.1 7.6 7.5 8.9

2.7 4.9 7.3 6.5 6.8 8.6 5.9 — 9.0 6.8 4.8 4.8 2.5 5.6

Ca Phos mg/dl mg/dl

Notes: Glu = glucose, BUN = blood urea nitrogen, Cr = creatinine, Bili = bilirubin, Chol = cholesterol, Trig = triglycerides, TP = total protein, Alb = albumin, Glob = globulin, Amy = amylase, Lip = lipase, AP = alkaline phosphatase, ALT = alanine aminotransferase, AST = aspartate aminotransferase, GGT = gamma glutamyl transpeptidase, CK = creatine phosphokinase, LDH = lactic dehydrogenase, Ca = calcium, Phos = phosphorus, N = normal individual in captivity.

6.5 2.5 1.3 1.3 2.0 2.4 2.2 3.0 4.6 2.0 2.6 1.2 1.5 1.2

132 131 119 170 232 140 167 314 135 232 252 172 207 131

1 2 3 4 5 6 7 8 9 10 11 12 13 N

62 44 44 41 44 74 56 108 57 44 47 84 40 40

Glu BUN Cr Bili Chol Trig mg/dl mg/dl mg/dl mg/dl mg/dl mg/dl

ID

TABLE 3 Serum Chemistry Findings in a Mass Stranding of False Killer Whales (Pseudorca crassidens)


Mass Strandings of Cetaceans 91


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Stranded short-finned pilot whales differed from the stranded false killer whales, in that no consistent biochemical or hematological abnormalities were present within the pod; however, individuals showed evidence of hemoconcentration, leukopenia, elevated serum creatinine, hyperbilirubinemia, hypocalcemia, and hypophosphatemia (Walsh et al., 1991). Similarly, in both strandings there was evidence of dehydration and stress that were supported by hemoconcentration and hyponatremia and by leukopenia, respectively. The hypocalcemia and hypophosphatemia were the result of unknown mechanisms but are not uncommon in stranded cetaceans, or subsequent to prolonged transport (Ewing, pers. comm.). Often, members of the pod have died by the time the rescue team intervenes. These animals should be necropsied to help determine what potential pathological processes are afflicting the pod. Sample collection is often difficult because of environmental conditions or logistics, but it is important that as thorough a necropsy as possible be performed (see Chapter 21, Necropsy). Table 4 illustrates the pathological findings for a group of stranded pilot whales from the Florida Keys in 1986 (Bossart et al., 1991). The pathological changes observed were diversified within the pod and even varied within individuals. The predominant findings were nonspecific gastrointestinal inflammation and degenerative changes. There was also marked lymphoid tissue depletion, suggesting chronic stress, immunosuppression, or cachexia (see Chapter 12, Immunology; Chapter 13, Stress). The histopathological changes were nonspecific although they were indicative of chronic progressive disease (Bossart et al., 1991). Based on blood work and necropsy results, it was evident that the animals involved in this stranding were not healthy at the time of intervention.

Disposition of Animals in a Mass Stranding After all animals have been tagged for identification and blood has been collected for clinical laboratory analyses, the rescue team must decide on the disposition of the animals in the group (see Figure 3). Because illness may be a major factor by the time a pod of whales strands, choices of what to do with the group may be complicated. It is important to consider two points. If illness is a major factor, a wide range of illness severity may be manifested within the group. Some individuals may be critically ill, whereas others may be only slightly debilitated. Second, there may be a combination of other factors, in addition to the illness, that determines where the whales strand. Geomagnetic field differences may help determine where an ill group is more likely to strand. Local storms, currents, tides, bottom topography, and environmental oddities may be contributing factors. Hours or days after being pushed back out to sea, the same animal may not be leading the group, or environmental factors may have changed; as a result, the group may not re-strand, but instead go back to sea, perhaps to die, and valuable information may be lost. With prior knowledge of illness within the group, it may be inappropriate simply to turn the pod out to sea. The choices available to the rescue team are dependent upon the size of the pod, background of the rescue team, environmental conditions, and the availability of rehabilitation facilities. Each stranding should be viewed as an individual event, with the initial goal being to learn as much as possible about the primary factors involved. For example, on the northeast coast of Cape Cod Bay, Massachusetts, there is an area where mass strandings of pilot whales regularly occur (Geraci et al., 1999). Blood results and histopathological findings do not entirely incriminate illness as the major stranding factor. It is suspected that the local coastline and the rapid tide changes are the primary factors contributing to these strandings, although morbillivirus has been found associated with numerous strandings since 1982 (Geraci et al., 1999).

a

N

+2(Pu) +2

+2(Pn)

+1(Pt) +3(Pn)

N

+2(Pt) +5 +2(Pn)

A (123 cm, M)

B (144 cm, F) C (292 cm, M)

D (323 cm, F)

E (328 cm, F) F (330 cm, F)

G (331 cm, F)

H (350 cm, F) I (380 cm, F) J (440 cm, M)

N +3 +5

N

N N

N

+5 +5

N

N +3 +3

N

N N

+1(Pn)

+2 N

+3

Pulmonary

Inflammation Intestinal

+2 +2 N

+2

N +1

+1

+3 +4

N

Cardiovascular

N +3 +3

N

+3 +3

N

N N

+3

Hepatic Degeneration

+5 +4 NE

NE

+5 +5

+5

+5 +5

+5

Lymphoid Depletion

+2 +3 NE

NE

+3 NE

N

N +5

+3

Adrenocortical Lipid Depletion

Kidney: pyelitis, necrotizing, chronic–active, multifocal, moderate — Subcutis: cellulitis, necrotizing, chronic–active, multifocal, severe Skeletal muscle: myositis, necrotizing, chronic–active, severe Skin: dermatititis, ulcerative, chronic–active, multifocal, severe — Pancreas: pancreatitis, fibrosing, chronic, multifocal, moderate Pancreas: pancreatitis, necrotizing, chronic–active, multifocal, moderate to severe Tumor: uterus, fibroleiomyoma — —

Other

Source: Bossart, G.D., Walsh, M.T., Odell, D.K., Lynch, J.D., Buesse, D.O., Friday, B., and Young, W.G., 1991, Histopathologic findings of a mass stranding of pilot whales (Globicephala macrorhynchus), Proceedings Second Marine Mammal Stranding Workshop, NOAA Technical Report.

b

Grade ranges (+1 = mild; +3 = moderate; +5 = severe). Animal identification indicates straight-line length in centimeters from tip of rostrum to fluke notch and sex (M = male, F = female). N = No specific lesions present; P = Lesions associated with parasites (n = nematode, t = trematode, c = cestode, u = unknown); NE = Not examined.

a

Gastric

Animal b ID (length, sex)

TABLE 4 Graded Histopathological Findings in a Mass Stranding of Pilot Whales (Globicephala macrorhynchus)


Mass Strandings of Cetaceans 93


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Euthanasia The realistic options facing a stranding response team must include the possibility of euthanasia. This procedure should never be implemented unless all other possibilities have been investigated and eliminated (see Chapter 32, Euthanasia).

Return to the Sea Rescue groups around the world differ in their reactions to mass strandings, with some limiting their response solely to returning the animals to the water. This solution, which assumes that all is well with both the individuals and the group as a whole, has met with mixed success (Odell et al., 1980). On the west coast of Florida, it is common for cetaceans that strand to be pushed back into the water and to re-strand, each time with increased mortality. Occasionally, the whales are never seen again, so some assume this is the best way to handle the problem. In strandings where health and/or illness have been investigated, this cannot be the sole response. While certain rescue groups feel they are doing the best thing for the pod, they are not considering that they are sending many or all of the whales out of sight to die. It should also be considered that, if some of the animals are infected with a fatal infectious disease, returning these animals to sea may result in further spread of the pathogen. In addition, a great amount of valuable information that could help in future strandings is lost when animals are prematurely released back out to sea. Disease problems affecting these groups may not be discovered or documented. Miniaturization of tracking devices has allowed transmitters to be temporarily applied to cetaceans (see Chapter 38, Tagging and Tracking), which should be considered a possible approach to study the survival of animals returned to the sea.

Survival of Treated Whales The approach to treatment of individuals from mass strandings is similar to that for any other marine mammal that is ill. Survival time of members of the two mass strandings mentioned earlier ranged from 2 days to 18 months. Because medical investigations into stranding events have been limited, it is not known what percentage of a pod of stranded whales may survive. It appears that the survival rate will be very low, with the chance of survival depending upon the stage of illness, the type of illness, and the adaptability and age of the individual. It must be assumed that survival of the pod will be low if members have already perished. A review of the treatments of nine stranded individuals that survived longer than 1 month indicated that most of these individuals continued to have recurrent bouts of illness. Premature release of these individuals may infect other healthy pods that would not have been exposed without human intervention. The recognition of the presence of infectious diseases in beached cetaceans has changed the approach to rehabilitation. Facilities with in-house collections that accept stranded animals put resident individuals at risk, unless all beached animals are placed in total isolation. Personnel working with beached animals must not have any contact with collection animals. Wet suits, food utensils, shower facilities, and handling equipment must be totally separate to eliminate vector transmission. Failure to implement full quarantine procedures can result in disaster (Bossart, 1995).

Conclusion To date, investigations into the causes of cetacean mass strandings have improved with the increased involvement and cooperation of oceanaria, rehabilitation facilities, academic institutions, and federal agencies. Increased financial support has increased the return of information, but more must be done to ensure the thoroughness of each investigation.



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Although histopathology and limited serology are becoming more common, it remains important to synthesize these data with the environmental, natural history, clinical, bacterial, toxic, and viral components to yield a comprehensive final evaluation of each stranding event. As new diagnostic tests are developed, retrospective analyses of archived tissues and serum are critical. To accomplish this goal, laboratories designated as receiving hubs for this material must be identified. It may be helpful to partner with colleagues in other countries who are already accomplished in specialized fields. This will require development of research gateways to allow easier passage of research material between experts. It must be remembered that the initiating factor(s) of a stranding may have occurred days or weeks before the animals encountered land, so that some strandings may not be explainable, even if all possible information is gathered. Only ongoing detailed examinations of mass strandings will slowly lead to understanding of this phenomenon.

Acknowledgments The authors thank the staff and participants in the Northeast and Southeastern U.S. Marine Mammal Stranding Networks, the National Marine Fisheries Service, Mote Marine Laboratory, Miami Seaquarium, and Dolphin Research Center for their involvement in the gathering of this information. They also thank Julia Zaias (University of Miami, Miami, FL) for editorial assistance, Teri Rowles for reviewing this chapter, and Jim Mead and the Marine Mammal Program at the Smithsonian Institution for their vigilance in the pursuit of information on cetaceans and for their compilation of information on mass strandings.

References Best, P.B., 1982, Whales, why do they strand? Afr. Wildl., 36: 6. Bossart, G.D., 1995, Morbillivirus infection: Implications for oceanaria marine mammal stranding programs, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CDROM Archives. Bossart, G.D., Walsh, M.T., Odell, D.K., Lynch, J.D., Buesse, D.O., Friday, R.B., and Young, W.G., 1991, Histopathologic findings of a mass stranding of pilot whales (Globicephala macrorhynchus), Proceedings Second Marine Mammal Stranding Workshop, NOAA Technical Report, 85–90. Bossart, G.D., Baden, D.G., Ewing, R.Y., Roberts, B., and Wright, S.D., 1998, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizootic: Gross, histologic, and immunohistochemical features, Toxicol. Pathol., 26: 276–282. Cetacean Distributional Database, Marine Mammal Program, Smithsonian Institution, Washington, D.C. Cordes, D.O., 1982, The causes of whale strandings, N.Z. J. Med., 30: 21. Dudok Van Heel, W.H., 1962, Sound and cetacea, Neth. J. Sea Res., 1: 402. Eaton, R.L., 1979, Speculations on strandings as burial, suicide, and interspecies communication, Carnivora, 2: 24. Frantzis, A., 1998, Does acoustic testing strand whales? Nature, 392(6671): 29. Fraser, F.C., 1934, Report on cetacea stranded on the British coast from 1927–1932, Br. Mus. Nat. Hist., 11. Fraser, F.C., 1946, Report on cetacea stranded on the British coast from 1933–1937, Br. Mus. Nat. Hist., 12. Fraser, F.C., 1953, Report on cetacea stranded on the British coast from 1938–1947, Br. Mus. Nat. Hist., 13. Fraser, F.C., 1956, Report on cetacea stranded on the British coast from 1948–1956, Br. Mus. Nat. Hist., 14. Geraci, J.R., 1978, The enigma of marine mammal strandings, Oceanus, 21: 38–47. Geraci, J.R., 1989, Clinical investigation of the 1987–88 mass mortality of bottlenose dolphins along the U.S. central and south Atlantic coast, Final Report National Marine Fisheries Service, U.S. Navy (Office of Naval Research), and Marine Mammal Commission, 63 pp.


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Geraci, J.R., and St. Aubin, D.J., 1977, Pathologic findings in a stranded herd of Atlantic white-sided dolphins, Lagenorhynchus acutus, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Geraci, J.R., and St. Aubin, D.J., 1979, Biology of marine mammals: Insights through strandings, U.S. Marine Mammal Commission, Report Number MMC-77/13, Washington, D.C., PB-293, 890. Geraci, J.R., Testaverde, S.A., Staubin, D.S., and Loop, T.H., 1976, A mass stranding of the Atlantic white-sided dolphin (Lagenorhynchus acutus): A study into pathobiology and life history, U.S. Marine Mammal Commission, Report Number MMC 75/12, Washington, D.C., PB-289, 361. Geraci, J.R., Anderson, D.M., Timperi, R.J., St. Aubin, D.J., Early, G.A., Prescott, J.H., and Mayo, C.A., 1989, Humpback whales (Megaptera novaeangliae) fatally poisoned by dinoflagellate toxin, Can. J. Fish. Aquat. Sci., 46: 1895–1898. Geraci, J.R., Harwood, J., and Lounsbury, V.J., 1999, Marine mammal die-offs, in Conservation and Management of Marine Mammals, Smithsonian Institution Press, Washington, D.C., 367–395. Kennedy, S., Smyth, J.A., Cush, P.F., McCullough, S.J., Allan, G.M., and McQuaid, S., 1988, Viral distemper now found in porpoises, Nature, 336: 21. Kirschvink, J.L., Dizon, A.E., and Westphal, J.A., 1985, Evidence from strandings for geomagnetic sensitivity in cetaceans, J. Exp. Biol., 120: 1–24. Klinowska, M., 1985a, Interpretation of the U.K. cetacean strandings records, Rep. Int. Whaling Comm., 35: 459. Klinowska, M., 1985b, Cetacean live stranding sites relate to geomagnetic topography, Aquat. Mammals, 11: 2–32. Klinowska, M., 1985c, Cetacean live stranding date relate to geomagnetic disturbances, Aquat. Mammals, 11: 109–119. Mead, J., 1997, Pathobiology of cetacean strandings along the Atlantic coast, 1976–1977, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Morimitsu, T., Nagai, T., Ida, M., Ishii, A., and Koono, M., 1986, Parasitogenic octavus neuropathy as a cause of mass stranding in odontoceti, J. Parasitol., 72: 469. Morimitsu, T., Nagai, T., Ida, M., Kawano, H., Naichuu, A., Koono, M., and Ishii, A., 1987, Mass stranding of odontoceti caused by parasitogenic eighth cranial neuropathy, J. Wildl. Dis., 23: 586–590. Odell, D.K., 1987, The mystery of marine mammal strandings, Cetus, 7: 2. Odell, D.K., Asper, E., Baucom, J., and Cornell, L., 1980, A recurrent mass stranding of false killer whales, Pseudorca crassidens, in Florida, Fish. Bull., 78: 171–177. Ridgway, S., and Dailey, M., 1972, Cerebral and cerebellar involvement of trematode parasites in dolphins and their possible role in stranding, J. Wildl. Dis., 8: 33–43. Sergeant, D.E., 1982, Mass strandings of toothed whales (Odontoceti) as a population phenomenon, Sci. Rep. Whale Res. Inst., 34: 1. Walsh, M.T., Beusse, D.O., Young, W.G., Lynch, J.D., Asper, E.D., and Odell, D.K., 1991, Medical findings in a mass stranding of pilot whales (Globicephala macrorhynchus) in Florida, Proceedings Second Marine Mammal Stranding Workshop, NOAA Technical Report 98, January, 75–83. Wareke, R., 1983, Whales, whale stranding—accident or design? Aust. Nat. Hist., 21: 4312. Wiley, D.N., Early, G., Mayo, C.A., and More, M.J., in review, The rescue and release of mass stranded cetaceans from beaches on Cape Cod, Massachusetts, USA: A review of some response action, Aquat. Mammals. Wilkinson, D.M., 1991, Report to the Assistant Administrator for Fisheries, in Program Review of the Marine Mammal Stranding Network, U.S. Department of Commerce, NOAA, NMFS, Silver Spring, MD, 171 pp. Wilkinson, D.M., 1996, National contingency plan for response to unusual marine mammal mortality events, U.S. Department of Commerce, NOAA Technical Memorandum, NMFS-OPR-9.


7 Careers in Marine Mammal Medicine Leslie A. Dierauf, Salvatore Frasca, Jr., and Ted Y. Mashima

Introduction All veterinarians working in the field of marine mammal medicine have many stories to tell about veterinary students and seasoned veterinarians with career changes in mind coming to them and asking for direction on where to find that perfect job in marine mammal medicine. One of the authors (S.F.) as director of education of the International Association for Aquatic Animal Medicine (IAAAM), for example, responds to an average of one to two e-mail inquiries per week from high school students, undergraduate and graduate students, veterinary students, or veterinary practitioners, regarding the availability of jobs in marine mammal medicine. Such a deceptively simple inquiry actually entails a long and complicated answer. Each individual career path represents a unique blend of what that person wants to do, what experience and training he or she brings to the pursuit, and what personal lifestyle choices that person wishes to honor (Dierauf, 1996). In 1994, the Society for Marine Mammalogy published a useful guide, which is available on the Web, that is the basis for some of the information in this chapter (Thomas and Odell, 1994). Other aspects come from the authors’ own personal searches for that “perfect job.” One may ask, “How can I have a great life, pursue my interest in marine mammals, and at the same time enthusiastically participate in this marvelous profession of veterinary medicine?” The choices really are very personal. Whether you are seeking a position in marine mammal clinical practice or marine mammal conservation and management, the opportunities available are varied and depend on your interests, skills, expertise, and abilities. One thing is certain: as a veterinarian with broad medical, scientific, and customer service expertise, you have excellent basic training in a variety of fields (Mashima, 1997), and can take your career in any direction that you wish. When you consider everything you are capable of doing, you will amaze yourself. One of the authors (L.A.D.) keeps this inspirational message on her desk, above a picture of a snow-covered, blue-skied mountain: “I am not in the habit of starting my day by thinking of things that I cannot get done!” Any one of the multitude of scientific, technical, and nontechnical topics/fields discussed in this textbook is a potential job opportunity for you.

Full-Time Employment Full-time jobs in clinical veterinary medicine of marine mammals are rare, and primarily limited to display facilities, the military, and rehabilitation centers. Currently in the United States, the authors estimate there are fewer than three dozen veterinarians employed in the fulltime practice of marine mammal medicine; a number of these are employed in marine research. 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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Even at the four Sea World facilities in the United States, where most “full-time” marine mammal veterinarians are employed, the caseload goes beyond marine mammals to birds, fish, and other marine organisms. Each year the selection criteria for positions in the field of marine mammal medicine become more stringent. In the opinion of one of the authors (S.F.), it is no longer a reasonable and wise career decision to consider yourself a viable candidate for these positions based solely on your classical veterinary education and degree. The competition for salaried positions and funding to perform clinically relevant research pertaining to marine mammals is intense. The viable candidate is someone who has developed skills in addition to formal veterinary training. These are skills in fields such as biomedical technology, computer science, population dynamics, public and environmental health, and conservation, which are complementary to formal veterinary medical training. Individuals with such skills often can improve their job opportunities, because they can present themselves as multifaceted professionals capable of multitasking at high levels and capable of filling more than one niche within the infrastructure.

Part-Time Employment Now that the concerns for full-time employment have been addressed, there are a number of ways to work as a marine mammal veterinarian on a part-time basis, either as a volunteer or consultant, in a variety of state and federal agencies, nonprofit private organizations, environmental groups, or in academia. In addition to clinical jobs, there are positions in marine mammal medicine involving preventive medicine, pathology, epidemiology, management, policy making, and public education, outreach, and awareness. More often than not, developing an expertise in some associated field, such as epidemiology, pathology, or education, may be a principal route into the field of marine mammal health management (King, 1996; Marshall, 1998; Smith, 1998a,b). The concept of conservation medicine can be well applied to marine mammal medicine. This movement blends conservation biology with veterinary and human medicine, and it is gaining rapid recognition as an interdisciplinary, team-oriented science (Jacobson et al., 1995; Aguilar and Mikota, 1996; Deem et al., 1999; Meffe, 1999; Society for Conservation Biology, 2000). Conservation medicine in the marine context addresses the application of biomedical principles and technology to global issues of ecology and environmental health. It also encompasses a wide range of interests, ranging from collaborative research in marine mammal population status to the effects of changes in marine ecosystems on marine mammal health and disease; from conservation efforts to protect vital habitats to concerns over international public health; from the effects of ecotourism to policy-making and funding opportunities for protection of natural resources and marine environments. Thus, although conventional clinical jobs may be few and far between, there are a myriad of opportunities that involve marine mammal health interests. You may create many of these opportunities, as you apply your background in alternative ways (Environmental Careers Organization, 1993; Gerson, 1996; Doyle, 1999; National Wildlife Federation, 2000).

Personality Traits and Other Tools Personality traits that lend themselves to exploration, risk-taking, and creativity are a plus in finding new career directions (Covey, 1990; Fassig, 1998; Johnson, 1998; Sylvester, 1998). Tools that come in handy are imagination, vigilance, practicality, patience, enthusiasm, and a willingness to dare to dream. These are the traits that lead to “making your own luck” (Wells, 1992). Luck is really the meeting of opportunity and preparation.


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Another set of tools that is vital to veterinarians who wish to move outside the traditional practice setting consists of those skills learned outside the profession itself. Such skills include creative writing, editing, networking, computer science, leadership, and critical thinking. Additional training or experience in conservation biology, ecology, population biology, environmental science, foreign languages, and journalism can help in developing these skills. Oftentimes, the result is a global perspective with big-picture views of the world, its people, and cultures, an awareness of the effects marine mammals and other animals have on our world, and the effects our world has on them. In reality, the majority of marine mammal medical skills will also be learned outside of formal veterinary training (Dierauf, 1996). These personal development strategies, professional improvement opportunities, and global perspectives are not improvement strategies unique to the field of marine mammal medicine. Some veterinary colleges have recognized the importance of these personal skills and the role that veterinary medicine can play in the realm of world health. They have developed didactic and active learning experiences in such fields as international veterinary medicine and population biology that address global concerns and apply the veterinary medical degree in alternative ways.

Summary Not everyone involved in marine mammal health is a veterinarian. Individuals who hold masters and doctorates in biomedical fields, such as molecular biology, cell biology, physiology, immunology, toxicology, neurobiology, ecology, and evolutionary biology, have contributed greatly to the advancement of marine mammal health over the past decades. Indeed, some of the most prolific and influential investigators in marine mammal biology have been nonveterinary professionals. The theme among all those individuals who have successfully developed careers in marine mammal health and medicine is excellence. Developing a reputation for excellence in some discipline and applying that excellence to the field of marine mammal health is the key to professional growth in this arena. In any case, this chapter is a generalized approach to identifying and seeking that “ideal” job, rather than an exacting formula for obtaining a position in marine mammal medicine. This chapter can be used as a guide, yet the decisions to be made are up to you alone. Use the suggestions in our “six-step method” as best suit your needs and desires for professional and personal development and fulfillment.

The Six-Step Method for Landing That Perfect Job Working with Marine Mammals 1. The First Step—Taking a Personal Self-Assessment The field of marine mammal medicine and conservation may look enchanting, but is it really for you? Do you have the personal desires and lifestyle needs that will fit into this professional field? What are your work ethics and interests? Will a job in the field of marine mammalogy fit your current time frame? Are there any particular patterns that have emerged in your career choices to date (Buss, 1998)? We would be remiss if we did not tell you that the field of marine mammal medicine today is less than lucrative in terms of salary and advancement. To date, the majority of vacancies have occurred in aquaria, academic institutions, and federal/state government agencies, because there are only so many coastal areas in the United States and abroad upon which to base a career in marine mammal medicine.


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First, you need to determine what exactly you are seeking with a position in the field of marine mammal medicine. Here are some questions to ask yourself, so that you have a clear picture of where you want to go in your professional career and what is most important to you in making any career change. We recommend that you not only read these questions, but also actually write down your answers to assist you as you move through the six-step process outlined in this chapter. Self-Assessment Questions Do you want to work part-time or full-time? Have you considered volunteering? Can you commit to an externship, internship, residency, or fellowship at this time? Is where you live important to you at this time? If so, where would you like to live? Do you have the means to live abroad, or are you planning to stay in the country where you currently reside? Do you have a family to support? If so, can you support your family in this career path? How motivated are you? Do you have the skills and training necessary for a position in this field? Do you have the time and resources available to take additional coursework or training? Does the position you are seeking fit your philosophy of life, lifestyle, and life goals? Are you ready to commit to a full-time job search, or are you peripherally interested at this time? Are you ready to commit to a job in a competitive field such as this? Have you paid enough attention to this field?

Have you taken time to work with a veterinarian in an aquarium or a teaching institution to appreciate the commitment of hours and effort that are required to maintain a job position? Do you realize that in some of the marine mammal medicine positions, especially in field research or clinical practice, the hours can be long, erratic, and unpredictable? If they involve administrative duties, these can entail daily paperwork, writing, reporting, and supervising. Because many marine mammal positions require you to be out of doors, even regular tasks and chores can become onerous if performed under extreme climatic conditions, such as scorching sun, brutal rain, unending wind, and rough seas. We urge you to consider each of these questions and issues seriously.

2. The Second Step—Categorizing Your Unique Skills, Strategies, and Approaches These days it appears that businesses, organizations, and institutions are searching for employees who stand out in a crowd. Tom Peters (1999) calls it “hiring to talent.” He frames whom to hire by looking for special “projects, passion, provocation, partnerships, politics, professionalism and performance.” He said that once, in pouring over 200 applications for a single position, he made his first cut by looking at the applications and watching for something peculiar; in this case, it was a computer scientist who had been entered into Ripley’s Believe It or Not for creating and baking a 1-ton cookie! A good foundation in small animal medicine and surgery and critical care medicine may serve you well in your marine mammal pursuits and casework. Some marine mammal clinicians have expressed to us that they prefer to hire individuals who have strong small animal medicine backgrounds and/or have completed small animal internships or residencies. In fields such as marine mammal medicine and conservation, potential employees exhibiting imagination and creativity often stand out from the rest. We believe it is scientifically founded, innovative



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thinking that will bring new marine mammal positions to the forefront, expanding the opportunities available for us all. Additional academic training and/or degrees can be helpful, as can short courses and continuing education in the field of marine mammal medicine. The ability to conduct self-motivated and self-motivating training and teaching and to participate in a volunteer capacity at facilities that cater to programs for advanced study and public involvement can add to your experiences in the field and build your professional skills and credentials. Volunteering in an organization, for no pay and hard work, can be an admission ticket to the world of paid employment, assuming you are productive and resourceful in choosing a particular role and how you focus on that role within the organization. For example, one of us (L.A.D.) came to be in charge of veterinary services (a paid position) at The Marine Mammal Center in Sausalito, California, by first volunteering every Sunday (over a year’s time) to set up a clinical laboratory and design a veterinary medical education course for the volunteers. However, in today’s economy, this may not be the most practical way individuals can acquire jobs in marine mammal health care. The advice often given by one of the authors (S.F.) regarding volunteerism is to strive to produce tangible results from your volunteer efforts and investments of time and expertise. This is especially true for students. Paid positions for veterinary students at display facilities or academic institutions are rare, and, when offered, the pay is often not commensurate with the effort. However, volunteer efforts may furnish opportunities to participate in clinical investigations or research projects that produce journal publications, conference presentations, or posters. Presenting your work at scientific conferences is an excellent way you can introduce yourself to large groups of potential employers or future collaborators. Some organizations, such as the IAAAM, encourage and support student presenters with competitions for student travel and conference presentation awards. On-the-job training, be it paid or unpaid, is always of value. Equal in importance to such active learning is discovering and committing to a mentor in the field (Harris, 1998). The mentor should be someone who can guide you and be an advocate for your career choices; someone who gives you an inside view of what the profession of marine mammal medicine and conservation is all about; someone who helps you build a base of contacts and networking individuals for future reference and support. All the authors have no doubts that the conscientious guidance and advice obtained from our mentors has been, and continues to be, integral to our career development. What tasks really fire you up? What tasks exhaust you? Richard Bolles (2000, p. 349) recommends that you make a list of all the things you enjoy doing with regard to work and play in general, and then categorize each item under one of these headings: “Skills with People,” “Skills with Things,” and “Skills with Information.” Bolles (2000, p. 79) also provides a list of 246 action verbs that describe a great variety of skills that, again, can be categorized under People, Things, and Data. How many of these action verbs relate to your skills and abilities? For example, are you a “people” person—Do you like mentoring, negotiating, instructing, supervising, persuading, speaking, serving, helping? Are you a “things” person—Do you like setting up things, working with precision instruments, operating technical devices, manipulating mechanical things, handling tools? Or are you a “data/information” person—Do you like collating, synthesizing, coordinating, analyzing, compiling, solving, computing, comparing? Once you have an idea of the variety of skills you have, write them down in order, beginning with the activities you enjoy doing the most. You will be surprised what clarity this simple exercise can bring to your marine mammal job search. This answers the question for you of “What do I want to do with marine mammals in my professional life?”


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3. The Third Step—Planning for Action and Timing The next step involves How to find the jobs that will give you the greatest satisfaction and opportunity to use your favorite skills. It is time to set some objectives and devise some options for planning your job-seeking strategy. Who knows you the best? It is likely to be your friends, family, peers, and colleagues. One of us (L.A.D.) “discovered” The Marine Mammal Center, when her friend, an art therapist, took her there during a summer afternoon outing. Ask the people who know you best to help you think of job opportunities and locate leads. With each lead, investigate the position and organization thoroughly to make certain each fits your current wants and needs. Use passive resources, such as telephone books, entertainment guides, the Internet, and on-line and hardcopy newsletters; sometimes these resources can trigger new job ideas, as well. Compare each job you come across with your prioritized list of skills and with your own strengths and weaknesses. Talk with anyone and everyone you meet who has the slightest connection to marine mammal medicine and health, to glean suggestions on other sources of information or other recommended organizations. Consider doing an elective “externship” that allows you to spend 4 to 6 weeks at a zoological park, aquarium, marine park, research facility, rehabilitation center, or government agency. After you find individuals who hold jobs you find attractive, ask them what they enjoy about their jobs, why they have kept their jobs, and how they obtained their jobs. Then make a list of the potential jobs and organizations and begin to investigate those people who are actually responsible for hiring to the kind of position you are seeking. Take a look at the section “Accessing Resources” at the end of this chapter, and the electronic job-hunting sources and ideas available in The Electronic Whale (Chapter 8), as well. In other words, it is just like school all over again; do your homework and you will succeed in gathering the information you need to make choices regarding the next phase of your professional career.

4. The Fourth Step—Making Choices The next step entails writing a job description for that perfect job, where you can use all your favorite skills, meet all your current lifestyle goals and objectives, and have some fun doing it. Try not to criticize or obstruct any ideas that might flow from your pen. Just keep writing, until you have on paper what your perfect job in the field of marine mammal medicine would be. This may seem like a fruitless, time-consuming exercise, but in reality it will truly clarify the direction you may want to take in choosing which positions to apply for, and then directing your career growth once you are in an organization. It will also insert some patience into your job search, recognizing that being in the right place at the right time may take time. You cannot really plan for the right time or the right place, but you can be prepared, and thereby recognize when the time and place are right. You will know. Now it is time to determine where you want to work. The best way to find where there are marine mammal medicine jobs is to network with people already in the marine mammal field. Choose one or more organizations you are interested in and start to nurture your networks. Find out what veterinarians or marine scientists already work there. Attend scientific marine mammal meetings, have coffee with these folks, get to know them, and, most importantly, let them begin to get to know you. Have patience, do not be overbearing, and make sure you ask the people you are networking with if they have time to talk with you. If they do not, ask them when (and where) would be more convenient. Be diplomatic and respectful of time in cultivating and maintaining your network. As another approach, if you are unable to make personal contact (although that is what these authors strongly recommend), pick up the telephone and call those facilities, organizations,



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institutions, or laboratories that most interest you. Visit the companies/institutions of your choice. Ask to see their lists of job openings and general job descriptions. If there are no written job descriptions, come prepared to ask a set of preapplication questions of the people responsible for hiring in the organizations of your choice. In the case of academic positions within the laboratory of a primary investigator, make every attempt to schedule a meeting and a tour with the investigator. Tour the campus and examine the locations of buildings and facilities that are likely to be important resources for you. Assess whether the support facilities are truly convenient and accessible when you evaluate the opportunity as a whole. Again, take your time, be mild-mannered, and do not waste/hog the time of the people who work at the facilities of interest to you. Prepare and carry with you one or more résumés that speak to the particular type of job or organizational framework of interest. If you see a job description that appeals to you, ask who is in charge of hiring for that position. Get the correct spelling of the person’s name, his or her title or position, and telephone number. Bring professional stationery and envelopes with you. Insert a made-to-order résumé and list of references in an envelope, hand-write a short note to the appropriate person, insert it in the envelope, and write the person’s name, title, and division or organization on the envelope. Ask the personnel office or the office assistant to hand-deliver this note for you. If there is an application form for the position, fill it out thoughtfully. Be neat, organized, and concise, providing the exact information the application seeks, no more, no less. In your answers, “lead with the lead”; begin with a sentence that directly answers the question the application asks. Mail or hand-deliver the application on time (or even prior to the closing or due date—do not fax an application or supporting documents or e-mail information, unless that is what the application asks for). Include a cover letter that tells the hiring person that you are very interested in the position and that urges that person to inspect your application in detail and seriously consider you as a candidate. Be patient. All things come to those who wait. One of the authors (L.A.D.) decided in 1977 that she wanted to go into the field of marine mammal medicine. Not until 1979 did she take a hands-on marine mammal medicine workshop and meet her mentors. Not until 1980 was she hired into a paying job at The Marine Mammal Center; it took another 10 years (1990) to move into the marine mammal policy and conservation medicine arena. On a regular and consistent basis, make friendly calls to the people with whom you have been networking, so they know that you continue to remain interested. Finally, remember that the early bird catches the worm; be persistent, resourceful, and friendly in your efforts and contacts.

5.

The Fifth Step—Preparing for the Interview

The hope is that your networking, homework, legwork, and follow-up calls and letters have brought you the opportunity for an interview. Never walk into an interview or respond to a phone call for an interview until you have prepared and composed yourself. Do not appear desperate (even if you feel that way!) or too eager (even if you are ecstatic) when you are contacted. Be calm, cool, collected, polite, professional—and ready! In the phone call inviting you to an interview, make sure you ask what type of interview format will be used: in-person, by telephone, one-on-one, small panel, large panel, tour through a number of different offices for a series of interviews, on-the-job, real-life situations, or a combi-nation of these formats. There are a number of questions (Ryan, 2000) you may want to ask yourself and answer in writing prior to any interview opportunity. So, as soon as you have any hint that you


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may be called for a telephone or in-person interview, begin preparing. Robin Ryan (2000) suggests that before you answer any preparatory questions, you first list as many as ten of your strongest traits. Then choose the five that most fit the job you have applied for, and rank those from 1 to 5. This is your five-point strategy. Consider these five points as your main answers to any of the questions posed here. Insert humor, enthusiasm, and anecdotes that demonstrate situations in which you successfully completed tasks related to the particular points you are presenting. Preparation is key; when someone tells you that you are lucky to be offered such an opportunity, be humble and recognize that you really do “make your own luck.” Tips for any interview (Dierauf, 1994): • The first 60 seconds of your interview are the most important; be prompt, neat in appearance, confident, and, above all, be prepared. Check your ego at the door. • Listen carefully to each question the interviewer asks you, pause, compose your thoughts, and then give an answer that is succinct, clear, and to the point. Use your five-point agenda whenever appropriate. Plan a number of different ways to deliver the same message. • Never take less than 20 or more than 90 seconds to answer a question. This ensures that the interviewer remains informed and energized by your presentations. • Remember that information and knowledge are power; the more you can absorb before your interview, the more smoothly the interview will proceed. Understand all aspects of any potential issue you may be asked to address. • If at all possible during the interview, do not discuss salary and benefits. This is a negotiation strategy you will want to work on if and when you are offered the job. This is just an interview. If the interviewer persists, ask what the salary range is for the position. Then deflect the question diplomatically by saying, “I believe the skills and experience I offer fit within that range,” or “That range is a bit lower than I had anticipated, but I am sure we can discuss that more fully at a later time, should you offer me this position.” • Have a rehearsed and practiced closing statement (60 seconds or less) to give yourself that final marketing sell before you exit the interview.

During an interview, you can anticipate being asked a number of standard questions. For example, the first things on any interviewer’s mind, although he or she may not express them out loud, are these two: Can you and will you do the job? Will you fit into the philosophy and mission of this organization/institution?

Work the answers to these often unasked questions into responses to actual questions, by talking about your current job and responsibilities, your commitment to your job, that you really find work enjoyable, and remember your five-point strategy. Assuming the person interviewing you is the person who will become your supervisor, answer in such a way that does not threaten that supervisor’s position in any way. You want to point out that you can complement his or her wishes and needs. Be sure that during the interview, if the interviewer is not clear or detailed enough, that you pleasantly ask for clarification or more detail. There are other common questions you should expect to be asked: Tell me about yourself (stick to your professional accomplishments, briefly summarizing your professional life over the past few years—keep it simple and short). Why should I hire you?



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What makes you think that you have the qualifications for this position? Why do you want this job? What are the features of your current job that you like the most? The least? Why did you leave your last job? Why are you unemployed? Why are there time gaps in your work history? What are your strengths? Your weaknesses?

All interviewers have their own reasons for asking the questions they do and in the manner they ask them. Prepare for unique questions or variations on them, such as: The Positive Approach—These are the interview questions that are the most enjoyable, where you can really shine, tell your stories and display your skills. Describe your current typical workday. Who was your favorite manager or supervisor and why? What do you know about this job and this organization? Name two or three things that are important to you in performing your job. What is the one thing you are proudest of in your (professional, not personal) life? What motivates you? What are you currently doing to improve yourself ? To you, what is the perfect job? The Negative Approach—Your responses to negative questions are best framed in a positive light. For example, take the question, give a brief answer, and then tell how you improved and/or learned from the situation, and how it made you grow and achieve greater success. Tell me about a time when you were criticized for poor performance. Describe a difficult co-worker. Tell me about one of your failures. How do you work under pressure? How do you handle stress? This job is a pressure-cooker. Can you handle it? Tell me something about your current boss that you dislike. Can you work odd hours, nights, weekends? Travel up to 20 days per month? How do you handle criticism? What was the most unpopular decision you ever made and what happened? What is the most difficult challenge you have ever faced (in your professional life)?

If the interviewer chooses such a negative approach, seriously consider whether you really want to work with this person. Was it a game he or she was playing, or is that person, with whom you will presumably be working, truly a negative sort? Regardless of the interviewer’s style, anticipate some not-so-common questions, such as the following, that you will definitely want to consider, to avoid being surprised and unprepared in your responses: What is the most recent book you have read? Who is the president/CEO of this company? Tell me about a personal goal you want to achieve. May I contact your current employer?

Also, be ready for any technical questions related to the scientific aspects of the job. The answers to the majority of these questions will be easy after all the homework and preparation you have done in the course of these first five steps. Be sure to write out your answers, so that you can review them prior to the actual interview.


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Anticipate that, at some time during your interview, the interviewer may ask if you have any additional questions. Prior to entering the interview, determine which of the following questions are appropriate for the marine mammal medicine position you are seeking: What is your professional background? What motivates you? Can you describe what my day-to-day responsibilities will be in this position? With whom will I be working? Tell me a little bit about their backgrounds and skills. Can you explain the organizational structure here? Describe the atmosphere and politics in this office. What financial and support resources underlie the department/program in which I will be working? Since coming to this organization and your current position, what would you describe as your two greatest successes? What do you feel are your greatest strengths? Weaknesses? What are your short- and long-term visions for this organization/institution? Do you anticipate hiring/firing staff in the next 24 months? For what reasons? What are the strengths and weaknesses of this organization? What is your management style and your favorite type of employee? Give me examples of three challenges that you and I can work together to resolve. I would like you to speak with my references. May we look at my reference list together?

Then close with what Ryan (2000) and Peters (1999) call the “Sixty Second Sell” or “Marketing the Brand YOU”—your own personal marketing ticket. Bring your interview back full circle by discussing what you do best, and how your enthusiasm and personality fit into, and complement, the mission and goals of the organization/institution, noting a few of your previous accomplishments that relate directly to the needs of the person hiring you and the job available. Be sure to tell the interviewer that, if you are hired, you intend to make a commitment to, and a difference in, the organization. Thank the interviewer, shake hands, smile, and calmly walk out. Go outside, sit down with pen and paper, and take notes about the interview and if you really believe you are a good fit for the job. Pat yourself on the back for a job well done. Follow up with a thank you note to the interviewer, and wait for the call.

6.

The Sixth Step—Starting Your New Job

In 1992, 24 scientists responded to a survey regarding career choices. From that survey, eight attributes important to any professional scientific career surfaced (Lebovsky, 1994): • • • • • • • •

Be knowledgeable in the subject of science; Develop and practice good communication skills; Be enthusiastic in the presentation of science; Support and encourage students and pre-professionals; Respect the abilities of students and peers and listen carefully to them; Be willing to give time and effort to help students; Relate subject matter to real-life situations; and Have compassion for, and commitment to, your profession.

How you communicate in your new career is very important. We are sure many of you already have excellent communication skills, and practice them every day, knowingly or unknowingly. Following is a basic list of communication tips one of us (L.A.D.) uses. These things are easy to do. The trick is to develop your own set of communication skills and practice



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using them every day. They will serve you well in your interactions with peers, colleagues, and hiring personnel in the field of marine mammal medicine. Communications Basics • • • • • • • • • • • • • •

Enhance and expand your oral and written communications (courses, practice, formal, informal). Train yourself to speak only after really listening and thinking. Do not let yourself get distracted when you are listening. Immerse yourself in a subject to learn it. Maintain a network of tried-and-true colleagues. Keep a positive attitude. Take nothing anyone says to you personally, even if it is so intended. Never take anything for granted. Steer away from viewing an issue as black or white, right or wrong. Take courses in teamwork, facilitation, mediation, and negotiation. Find a clear window of time (at least two 15-minutes periods) to think every day. Work at developing multiple options. Take risks; embracing risk is an exciting and energizing challenge. Have fun and keep your sense of humor.

Accessing Resources Resources are what the majority of your efforts will revolve around as you plan your strategies and needs for a career in marine mammal medicine. First, we invite you to consider contacting marine mammal specialists who have contributed to this edition of the Handbook of Marine Mammal Medicine as sources of career information and ideas. In addition, the majority of programs, organizations, and other information sources listed below with their Web site addresses can provide greater detail, including contact information. The Electronic Whale (Chapter 8) provides further sources of electronic information. The following list of professional resources is not intended to be exhaustive. Opportunities listed below may change in terms of content, instructors, requirements, and/or dynamics. It is the responsibility of self-motivated individuals to investigate the current status of opportunities that interest them.

Internships and Residencies Matched Internships Kansas State University, College of Veterinary Medicine, Manhattan, KS http://www.vet.ksu.edu The Ohio State University, College of Veterinary Medicine, Columbus, OH http://www.vet.ohio-state.edu University of Georgia, College of Veterinary Medicine, Athens, GA http://www.vet.uga.edu University of Michigan, College of Veterinary Medicine (also residencies), East Lansing, MI http://www.cvm.ms.edu


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These matched internship programs concentrate to varying degrees on exotic, wildlife, and zoo animals in the format of a rotating 1-year internship through a veterinary teaching hospital. These programs are not aquatic specific, but each offers open rotations and vacation in which to accomplish aquatic studies. Each of these programs participates in the Veterinary Medical Intern-Resident Matching Program, administered through the American Association of Veterinary Clinicians. http://cvm.msu.edu/~judy/aavcl.htm Matched Residencies North Carolina State University, College of Veterinary Medicine, Raleigh, NC http://www.cvm.ncsu.edu University of California, Davis, School of Veterinary Medicine, Davis, CA http://www.vetmed.ucdavis.edu University of Florida, College of Veterinary Medicine, Gainesville, FL http://www.vetmed.ufl.edu University of Tennessee, College of Veterinary Medicine, Knoxville, TN http://web.utk.edu/~vetmed/default.html University of Wisconsin, School of Veterinary Medicine, Madison, WI http://www.vetmed.wisc.edu

Each of these matched residency programs concentrates on exotic, wildlife, aquatic, and zoo animals in the context of a multiyear residency program through a veterinary teaching hospital and participates in the Veterinary Medical Intern-Resident Matching Program, administered through the American Association of Veterinary Clinicians. Individuals interested in residencies should contact the colleges offering such programs for admission requirements and application policies, and to introduce themselves to instructors. In addition, the dynamics of such programs may vary with regard to affiliations with regional aquariums and zoos. Other Internships

Internships at aquaria or rehabilitation centers: Mystic Aquarium, Mystic, CT http://www.mysticaquarium.org National Aquarium at Baltimore, Baltimore, MD http://www.aqua.org New England Aquarium, Boston, MA http://www.neaq.org SeaWorld, San Diego, CA http://www.seaworld.com The Marine Mammal Center, Sausalito, CA http://www.tmmc.org

These are veterinary internships, which are oriented to aquatic animal, for periods of 1 year or less, by arrangement, and are offered by institutions that are independent of the Veterinary Medical Intern-Resident Matching Program. The application policies and terms are determined by the admissions committee of each particular institution, and the content and experiences offered vary with the collection of animals being maintained, the research and veterinary services offered, and the affiliations established with other academic or research



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institutions. It is advisable to contact these institutions directly to learn of their unique application policies. Internships at zoos with exposure to aquatic animal medicine: Birmingham Zoo, Birmingham, AL http://www.birminghamzoo.com Brookfield Zoo, Chicago, IL http://www.brookfieldzoo.org Columbus Zoo, Columbus, OH http://www.colszoo.org Louisville Zoological Gardens, Louisville, KY http://www.iglou.com/louzoo John G. Shedd Aquarium and Lincoln Park Zoo, Chicago, IL http://www.shednet.org and http://www.lpzoo.com Smithsonian National Zoological Park, Washington, D.C. http://natzoo.si.edu St. Louis Zoo, St. Louis, MO http://www.stlzoo.org

These are veterinary internships offered by institutions independent of veterinary teaching hospitals, although most collaborate with regional research institutions and/or veterinary colleges. The conditions for application vary. It is advisable to contact these institutions directly to inquire about their programs. Internships affiliated with institutions or agencies: Alaska SeaLife Center, Seward, AK http://www.alaskasealife.org California Department of Fish and Game/UC Davis Wildlife Health Center, Davis, CA http://www.vetmed.ucdavis.edu/whc The Smithsonian Institution, Conservation and Research Center, Front Royal, VA http://www.si.edu/crc University of Alabama, Dauphin Island Sea Lab, Marine Sciences Program, Dauphin Island, AL http://www.disl.org The Wildlife Center of Virginia, Waynesboro, VA http://www.wildlifecenter.org

Graduate Degree Programs Programs with aquatic and marine mammal emphasis (from departments outside veterinary schools) Department of Biology, San Francisco State University, San Francisco, CA http://www.sfsu.edu/~biology Department of Biology, University of Alaska Southeast, Juneau, AK http://www.jun.alaska.edu


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Department of Pathobiology and Veterinary Sciences, University of Connecticut, Storrs, CT http://www.lib.uconn.edu/CANR/patho/index.html Department of Zoology, College of Biological Science, Guelph, Ontario, Canada http://www.uoguelph.ca/graduate studies Aquatic Pathobiology Center, Department of Pathology, School of Medicine, University of Maryland, Baltimore, MD http://som1.umaryland.edu/aquaticpath/ Aquatic Animal Disease Research and Diagnostic Center, School of Marine Science, The Virginia Institute of Marine Science, Gloucester Point, VA http://www.vims.edu/

These programs are graduate degree programs (i.e., Master’s and Ph.D.) offered by university departments or schools with faculty expertise in aquatic animal health. They are independent of veterinary teaching hospitals, although some, such as the Department of Pathobiology and Veterinary Sciences at the University of Connecticut, educate veterinarians in specialty training programs (e.g., veterinary anatomical pathology). The faculty of these programs determines the program offerings, and application policies vary according to the institution. This list of degree programs is not exhaustive; other programs are available and equally worthwhile. Interested individuals should investigate the course offerings and research opportunities at these and other institutions for programs that match their interests. Alternative sources of career opportunities include the Web sites, journal publications, and newsletters of following organizations: the American Association of Zoo Veterinarians, the Alliance of Veterinarians for the Environment, the American Veterinary Medical Association, the American Association of Zoos and Aquaria, the American Association of Wildlife Veterinarians, the Wildlife Disease Association, and the International Association for Aquatic Animal Medicine (see Chapter 8, The Electronic Whale).

Other Related Programs American Veterinary Medical Association, Government Relations Division, Schaumburg, IL and Washington, D.C. http://www.avma.org Center for Coastal Studies, Provincetown, MA http://www.coastalstudies.org Center for Marine Conservation, Washington, D.C. http://www.cmc-ocean.org Center for Oceanic Research and Education, Essex, MA http://www.coreresearch.org Conference on Trade in Endangered Species, U.S. Fish and Wildlife Service, Washington, D.C. http://international.fws.gov Dolphin Internship Program, Honolulu, HI http://www.pacificwhale.org/internships Global Green, USA, Green Cross International, Washington, D.C. http://www.globalgreen.org Long Island University Southampton Campus College of Marine Science, Southampton, NY http://www.southampton.liu.edu



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Moss Landing Marine Laboratories, Moss Landing, CA http://www.mlml.calstate.edu Oregon State University School of Oceanography, Newport, OR http://www.oce.orst.edu PAWS Wildlife Center, Lynnwood, WA http://www.paws.org/wildlife Scripps Research Institute, La Jolla, CA http:/www.scripps.edu SeaWorld, Orlando, FL; San Diego, CA; San Antonio, TX; Aurora, OH http://www.seaworld.com Stanford University Hopkins Marine Station of Behavior, Pacific Grove, CA http://www-marine.stanford.edu Texas A&M University, Galveston, TX http://www.marinebiology.edu University of Alaska College of Fisheries and Ocean Sciences, Fairbanks, AK http://www.uaf.edu University of Alaska Southeast Department of Marine Biology, Juneau, AK http://www.uas.alaska.edu University of California Long Marine Laboratory, Santa Cruz, CA http://www.ganesa.com/ecotopia/long.html University of Hawaii Marine Option Program, Honolulu, HI http://www.uhhmop.hawaii.edu University of Washington, College of Ocean and Fishery Sciences, Seattle, WA http://www.cofs.washington.edu Wildlife Conservation Society, Bronx, NY http://www.wcs.org Woods Hole Oceanographic Institute, Falmouth, MA http://www.whoi.edu

Although less widely publicized and broader in scope than medicine alone, these programs relate to marine mammals, marine sciences, and marine research, policy, and/or environmental advocacy.

Advanced Training Programs AQUAMED, An aquatic animal pathobiology course, sponsored by the Gulf States Consortium of Colleges of Veterinary Medicine at Auburn University, Mississippi State University, Louisiana State University, Texas A&M University, and the University of Florida; presented at the Louisiana State University School of Veterinary Medicine, Baton Rouge, LA http://www.vetmed.lsu.edu/aquamed.htm AQUAVET, A program in aquatic veterinary medicine, sponsored by the School of Veterinary Medicine at the University of Pennsylvania and the College of Veterinary Medicine at Cornell University; presented in collaboration with the Marine Biological Laboratory, the Northeast Fisheries Science


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Center of the National Marine Fisheries Service, and Woods Hole Oceanographic Institute, Falmouth, MA http://zoo.vet.cornell.edu/public/aquavet/aquavet.htm ENVIROVET, An intensive short course in wildlife and ecosystem health in a developed country and an international development context, sponsored by the College of Veterinary Medicine, University of Illinois at Urbana-Champaign, IL http://www.cvm.uiuc.edu/vb/envirovet/ MARVET, An intensive short summer course in marine mammal medicine presented by Dr. Raymond Tarpley at Texas A&M [email protected]

Fellowships American Association for the Advancement of Science, Washington, D.C. http://www.aaas.org American Veterinary Medical Association Congressional Science Fellowships, Washington, D.C. http://www.avma.org/avmf/csfapp.htm David H. Smith Conservation Research Fellowship Program http://consci.tnc.org/Smith.htm Harbor Branch Oceanographic Institute, Fort Pierce, FL http://www.hboi.edu International Oceanographic Foundation, Miami, FL http://www.rsmas.miami.edu/divs/mbf Sea Grant College Programs, Sea Grant Colleges and Universities nationwide (U.S.) search the web for Sea Grant College Fellowships

Scientific Societies and Membership Organizations Alliance of Veterinarians for the Environment http://www.AVEweb.org American Association of Wildlife Veterinarians http://www.aawv.net American Association of Zoo Veterinarians http://www.worldzoo.org/aazv/aazv.htm American Cetacean Society http://www.acsonline.org American College of Zoological Medicine http://www.worldzoo.org/aczm American Veterinary Medical Association http://www.avma.org American Zoo and Aquarium Association http://www.aza.org European Association for Aquatic Mammals http://www.eaam.org



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International Association for Aquatic Animal Medicine http://www.iaaam.org International Society for Ecosystem Health http://www.oac.uoguelph.ca/ISEH/index.htm National Sea Grant Program http://www.nsgo.seagrant.org Sarasota (FL) Dolphin Research Program http://www.mote.org/~rwells Society for Conservation Biology http://conbio.rice.edu/scb Student Conservation Association http://www.sca-inc.org The Society for Marine Mammalogy http://pegasus.cc.ucf.edu/~smm/about.htm Wildlife Conservation Society http://wildlifedisease.org Women’s Aquatic Network http://orgs.women.connect.com/WAN/welcome.html World Veterinary Association http://www.worldvet.org

One additional Web site offers a large array of additional marine mammal Web resources: http://ourworld.compuserve.com/homepages/jaap/mmmain.htm

Many of the resource organizations listed in this chapter maintain directories of their members by state to use for contact and networking purposes. They also produce newsletters and hold regular conferences and training workshops, which often involve roundtables on careers in marine mammal sciences and medicine (see Chapter 8, The Electronic Whale, for additional references related to marine mammal medicine).

Recommendations and Conclusions Although this chapter offers no guarantees for finding a position in marine mammal medicine, if you follow the general recommendations, the six-step method, and access the information resources, as well as remember the six recommendations below, you will make your own luck and may actually find that perfect job in marine mammal medicine or conservation. • • • • • •

Keep your eyes and ears open and keep networking. Be opportunistic. Find a mentor and work with that person as often as possible. Be patient. Maintain a public or professional presence. Be persistent.


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Acknowledgments The authors thank Scott Newman and Gwen Griffith for peer-reviewing this chapter, and especially Jocelyn Catalla for her Web research and for her perspectives from the point of view of a student. In addition, the authors thank the members of the MarMam and Wildlife Health listserves for responding so enthusiastically to our listserve question: “What are your favorite marine mammal Web sites?”

References Aguilar, R.F., and Mikota, S.K., 1996, To reach beyond: A North American perspective on conservation outreach, J. Zoo Wildl. Med., 27(3): 301–302. Bolles, R.N., 2000, What Color Is Your Parachute, 2000, Ten Speed Press, Berkeley, CA. Buss, D.D., 1998, Career development pathways in veterinary medicine, Convention notes, American Veterinary Medical Association, 135th Annual Convention, July 25–29: 114–115. Covey, S.R., 1990, Seven Habits of Highly Effective People, Covey Leadership Center, Provo, UT, 6 audiotapes. Deem, S.L., Cook, R.A., and Karesh, W.B., 1999, International opportunities in conservation medicine, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 860–862. Dierauf, L.A., 1994, Potomac fever: I had it bad! in From the Lab to the Hill: Essays Celebrating 20 Years of Congressional Science and Engineering Fellows, Fainberg, A. (Ed.), American Association for the Advancement of Science, Washington, D.C., 31–35. Dierauf, L.A., 1996, The Career Changing Tool Kit, Connections Newsl. Alliance Vet. Environ., 1(1): 4–5. Doyle, K. (Ed.), 1999, The Complete Guide to Environmental Careers in the 21st Century, Island Press, Washington, D.C., 447 pp. Environmental Careers Organization, 1993, The New Complete Guide to Environmental Careers, Island Press, Washington, D.C., 364 pp. Fassig, S.M., 1998, Job-seeking skills, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 753–755. Gerson, R., 1996, How to Create the Job You Want: Six Steps to a Fulfilling Career, Enrichment Enterprises, Austin, TX, 201 pp. Harris, J.M., 1998, Leo K. Bustad, DVM, Ph.D.: A veterinarian for all seasons, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 449–450. Jacobson, S.K., Vaughan, E., and Miller, S.W., 1995, New directions in conservation biology: Graduate programs, Conserv. Biol., 9(1): 5–17. Johnson, S., 1998, Who Moved My Cheese? G.P. Putnam’s Sons, New York, 94 pp. King, L.J., 1996, Seven habits of highly effective globalized veterinarians, J. Vet. Med. Educ., Winter: 45. Lebovsky, A., 1994, The role of college and precollege science teachers in determining the education and career choices of Congressional fellows: A legacy of the class of 1990–1991, in From the Lab to the Hill: Essays Celebrating 20 Years of Congressional Science and Engineering Fellows, Fainberg, A. (Ed.), American Association for the Advancement of Science, Washington, D.C., 383–386. Marshall, K.E., 1998, Twenty laws of successful job hunting in the veterinary jungle, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 758–760. Mashima, T.Y., 1997, Conservation and Environmental Career Opportunities, Connections Newsl. Alliance Vet. Environ., 2(1): 2–3. Meffe, G.K., 1999, Conservation medicine, Conserv. Biol., 13: 953–954. National Wildlife Federation, 2000, The 2000 Conservation Directory: A Guide to Worldwide Environmental Organizations, 45th ed., Washington, D.C., 544 pp. Peters, T., 1999, Reinventing Work: Fifty Ways to Transform Every Task into a Project That Matters, Alfred A. Knopf, New York, 28 pp.



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Ryan, R., 2000, Sixty Seconds and You’re Hired, Penguin Books, New York, 175 pp. Smith, C.A., 1998a, How students and practitioners can prepare for international opportunities, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 863–865. Smith, C.A., 1998b, Career Choices for Veterinarians: Beyond Private Practice, Smith Veterinary Services, Leavenworth, WA, 255 pp (see http://www.smithvet.com). Society for Conservation Biology, 2000, Symposium 7 on Conservation Medicine: The ecological context of health, 14th Annual SCB Meeting, Program and Abstracts, Missoula, MT, June 9–12: 102. Sylvester, N., 1998, Leadership skills for the new millennium: Interpersonal skills, Convention notes, American Veterinary Medical Association, 136th Annual Convention, July 10–14: 772–775. Thomas, J., and Odell, D., 1994, Strategies for pursuing a career in marine mammal science, Suppl. Mar. Mammal Sci., 10(2), April, The Society for Marine Mammalogy, Allen Press, Lawrence, KS, 14 pp. Wells, W.G., Jr., 1992, Working with Congress: A Practical Guide for Scientists and Engineers, American Association for the Advancement of Science, Washington, D.C., 153 pp.



8 The Electronic Whale Leslie A. Dierauf

Introduction On January 1, 2000, an Alta Vista search engine Web search for “marine mammal medicine” yielded 1,196,440 matches! Just prior to sending the chapters for this textbook off to the publisher, a second search was conducted using the same search phrase and again on Alta Vista; this time we found 11,426,338 matches, a tenfold increase in sites in less than 1 year! We also asked a number of listserves what were their members’ favorite Web sites pertaining to marine mammal medicine; we received over 50 responses from people around the world, many of whose suggestions are noted in this chapter and in Chapter 7 (Careers). These kinds of numbers provide but a hint of the explosion of Internet-based information that is occurring. Accessing information and products on the Internet is the wave of the future, and the future is here today.

Using Your Head on the Web Along with the World Wide Web to access information has come a tangle of difficulties. Reading materials on the Web really is no different from scientifically reviewing a potential paper for publication in a scientific journal. First, you must scrutinize the document and its authors to determine if the paper is even worthy of consideration. Then, using your best scientific judgment, you must decide if what you are reading is valid. The Web has no quality control per se; anyone in the world can represent him or herself as a marine mammal expert. Peer review is often lacking. Web writers span the spectrum from a leading expert in the field, who includes superb references and acknowledgments of peer reviewers, to someone with primarily an emotional interest in marine mammals, with minimal factual information and few to no scientific citations to back up assumptions or conclusions. We must each ensure that the marine mammal medicine and conservation information that comes online is accurate, scientifically based, and statistically valid. Since the public will have access to any scientific information online, electronic publications will need to be written in plain language, so that we, as veterinarians, communicate our scientific information to the public in an understandable and comprehensible fashion, just as if we were in an examination room trying to explain a disease process to a pet owner. Electronic information can be unbiased scientific results, or it can be advertisements for products, goods, or services of commercial ventures. Simply reading raw data can lead the information gatherer to misleading and incorrect conclusions. Accessing electronic information can be stressful. Try as we might, we expend more paper now in printing out the information we need than

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we did prior to the electronic age. Perhaps this too will change as time progresses. Perhaps in the future, Web sites of all kinds will have internal search methodologies that will allow a viewer to print specific sections of an article, and search more easily and quickly for specific detailed information, rather than getting ensnarled in the Web site. We look to a future in Internet technology where we all have the skills to know how best to frame a medical question, to use appropriate and accurate databases to access that information, to apply the answer to our marine mammal work, and, even more importantly, to be able to do this on-site in the field, just like a poolside rapid diagnostic test. So, you are urged to use your “sixth sense,” and if you have any doubts about what you are reading on a Web page, please be certain to check with known experts in the field before utilizing any potential diagnoses and the techniques and/or treatments the Web article recommends.

Reference Databases General Biomedical and Veterinary Medical Sites Conducting searches of the scientific literature by traditional methods, such as a library search, can be time-consuming, tedious, and expensive. Once you find the article you need, if it is in the library at all, you then need to photocopy it and carry it home to read. With each passing year, however, online searchable scientific reference databases become more numerous, more helpful, and more easily browsed. Following is a list of those most applicable online reference sources for accessing biomedical, veterinary medical, and/or marine mammal medical literature. The University of Michigan School of Information and Library Studies manages a series of Internet resource guides covering a huge number of subjects, one of which is veterinary medicine: http://www.lib.umich.edu/chhome.html

Michigan also has an electronic library that provides reliable access to scientific Internet resources. The site listed here allows you to enter the science and environment collection: http://mel.lib.mi.us

The San Diego Library Consortium is a searchable database by author, subject, title, or biomedical subject, and links to other California state system universities, so it is quite complete. Access it at: http://circuit.sdsu.edu

The U.S. Department of Agriculture, Food and Drug Administration maintains a database of biological collections on the Internet. The database covers specific subject matter and a large array of journals, which can be accessed: http://vm.cfscan.fda.gov/~frf/biologic.html

ProMED is a scientific information request site, on which animal science papers can be located. Although designed for physicians, this site contains invaluable diagnostic and therapeutic information and, therefore, can be useful in marine mammal clinical practice: http://promed-windows.com

Grateful Med and PubMed through the U.S. National Library of Medicine homepage is your entry to searches of Medline, standard medical vocabulary, public health, general medical, veterinary medical, and scientific literature abstracts, catalogs, databases, and disease research. These databases give you access to more than 20 billion scientific citations and abstracts, and cover French, Spanish, Portuguese, and Russian biomedical literature, in addition to English. The National Cancer Institute has similar online access to biomedical topics and literature.


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U.S. National Library of Medicine http://www.nlm.nih.gov Grateful Med http://igm.nlm.nih.gov/igm_intro/title.html PubMed http://www.nlm.nih.gov/pubs/factsheets/pubmed.html National Cancer Institute http://www-library.ncifcrf.gov

SNOMED (Systemized Nomenclature of Human and Veterinary Medicine) is a conceptbased reference site related to record keeping, laboratory and clinical pathology system tracking, decision-support systems, disease registries, and more. It also identifies and defines veterinary medical standardized terminology, rather like a veterinary medical dictionary (Monti, 2000): http://www.snomed.org

The U.S. Fish and Wildlife Service, National Conservation Training Center (NCTC) in Shepherdstown, West Virginia has an online conservation library. Articles, journals, and scientific literature related to a multitude of conservation issues can be searched by accessing: http://training.fws.gov/library

NetVet is an ingenious site (also accessible through the AVMA Web site) developed in 1993 by a veterinarian now at Washington University in St. Louis, Missouri. The site contains a wealth of information about veterinary medical and animal resources available on the Internet; it references hundreds of veterinary and animal health–related Web sites through its Electronic Zoo, and is updated regularly. In 1995 alone, more than 650,000 computer users referenced this site. Within NetVet is a general reference site for writers, which includes dictionaries, encyclopedias, virtual libraries, and other valuable resources you may need if you are writing scientific or lay literature on marine mammals. Be sure to contact the NetVet site and have your new domains included on the Electronic Zoo list. The NetVet site can give your scientific publications excellent public and scientific exposure. American Veterinary Medical Association http://www.avma.org NetVet http://netvet.wustl.edu NetVet specific to Marine Mammal Information http://netvet.wustl.edu/marine.htm

Model Web Sites and Evidence-Based Medicine The Health on the Net (HON) Foundation in Switzerland is a nonprofit organization intent on demonstrating the benefits of the Internet and related technologies to the fields of medicine and health care. Available in both English and French, HON includes Web site listings, journal articles, multimedia, and health news to provide integrated search results. The Organized Medical Networked Information (OMNI) is the self-described “United Kingdom’s gateway to high quality biomedical Internet resources.” OMNI relies on “unbiased, high quality, internetbased resources relevant to the medical, biomedical, and health communities.” These model Web sites insist that medical information on the Internet be peer-reviewed and “given [only] by medically trained and qualified professionals” (HON). Both sites welcome relevant resource


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additions. The veterinary profession would do well to model its information for the Internet following the guidelines of these organizations and to utilize the opportunity to add its scientific works to these databases. Health on the Net http://www.hon.ch Organized Medical Networked Information http://omni.ac.uk

Evidence-based medicine has a number of health and human medicine guidelines (AAFP, 1999), which the marine mammal medicine community would be wise to follow. For use in your scientific writing, the author recommends the following sites: University of Washington Library http://www.hslib.washington.edu/clinical/guidelines.html U.S. Government Guidelines http://www.guideline.gov

Marine Mammal–Related Listserves One of the more rapid ways to gather information is through a listserve. A listserve is a mail system for creating, managing, and controlling electronic mailing lists of names and addresses. Messages, questions, answers, and announcements are sent to groups of people with similar interests. You can subscribe to and unsubscribe from a listserve as your time and commitment warrant. The two listserve sources marine mammal scientists use most commonly are MarMam and WildlifeHealth. To subscribe to the MarMam listserve, send an e-mail message to: [email protected]

For the WildlifeHealth listserve, send an e-mail message to: [email protected]

You can join these listserves by typing in the Web address, then in the body of the e-mail inserting “subscribe” “marmam” or “wildlifehealth” followed by “Yourfirstname Yourlastname” on the subject line, and sending it electronically. To post messages, use: [email protected] and [email protected]

To contact the editors for MarMam, e-mail: [email protected]

To contact WildlifeHealth within the Wildlife Information Network in the United Kingdom, e-mail: [email protected]

MarMam—Marine Mammal Conservation and Discussion—list functions as an exchangeof-ideas location. The types of messages posted at MarMam range from requests for information to case studies to announcements of meetings and training opportunities to book reviews and journal abstracts. The WildlifeHealth listserve, originally set up through the National Wildlife Health Center (NWHC), which is a science center within the Biological Resources Division of the U.S. Geological Survey in Madison, Wisconsin, addresses wildlife health and



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facilitates the exchange of questions, answers, general information, case histories, and other concerns regarding wildlife health; any member of the listserve can post information, questions, answers, or concerns at the site. Both sites offer free access and unlimited use. Each site is archived, so past messages can be viewed and retrieved.

Other Internet Discussion and Marine Mammal Information Lists There are currently at least four major information sites where e-mail discussion groups, chat rooms, announcements, and information lists can be registered, advertised, and accessed, including Lyris (Lyris Technologies, Inc., Berkeley, CA), Majordomo (Great Circle Associates, Mountain View, CA), LISTSERV (L-Soft, Landover, MD), and ListProc (Corporation for Research and Educational Networking (CREN, Washington, D.C.). Lyris http://www.lyris.net Majordomo http://www.greatcircle.com/majordomo [shareware] LISTSERV http://www.listserv.net CREN http://www.listproc.net [for UNIX users] List Identification http://tile.net/lists

The sites listed here are excellent linkage points for marine mammal medicine and science sites. Dalhousie University http://is.dal.ca/~whitelab/links.htm Five Colleges Coastal & Marine Sciences http://www.science.smith.edu/departments/marine Marine Mammal Net http://marinemammal.net National Marine Mammal Laboratory http://nmml.afsc.noaa.gov/library/resources/resources.htm Whale Net http://whale.wheelock.edu

Online Marine Mammal Journals and Textbooks In this age of electronic information, many veterinary medical journals, including marine mammal journals, are online, and textbooks are expected to be online soon. If you are an electronic textbook editor, ensure that your authors electronically submit their publications only through a quality-control gateway, and only after peer review. Materials with highquality electronic information will serve the public well, will improve accessibility, and will lead to lower costs for accessing information and greater opportunity for interacting electronically with colleagues regarding marine mammal medical information. This is already happening on a regular basis in the medical profession (BioMedicina, 1999) and at academic institutions. However, even in the medical profession, not enough physicians


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have the skills and abilities that are required to frame diagnostic queries or clinical questions or to use the databases available to locate and apply the answers to the care of their patients. The author urges marine mammal veterinary medical specialists to participate in this arena of high-quality and quality-controlled electronic information. The journal Marine Mammal Science is available online, as are additional journal and textbook reference materials. Marine Mammal Science online http://pegasus.cc.ucf.edu/~smm/mms.htm Library of Michigan http://mel.lib.mi.us/science/auth.html National Council for Science and the Environment http://www.cnie.org/journal.htm Nova Southeastern University, Ocean Center Library http://www.nova.edu/cwis/oceanography/library.html San Diego State University http://circuit.sdsu.edu University of Buffalo Science and Engineering Library http://ublib.buffalo.edu/libraries/units/sel/collections/ejournal2.html#a University of Montreal Beluga Whale Info http://www.medvet.umontreal.ca/services/beluga/index_an.html U.S. Fish and Wildlife Service Literature Search http://training.fws.gov/library

Fellowships, Foundations, and Grants Fellowships

Congressional Science Fellowships are paid positions, sponsored by the American Veterinary Medical Foundation (AVMF) and the American Association for the Advancement of Science (AAAS). They are awarded competitively to scientists, who serve for 1 year in Washington, D.C., for either the U.S. House of Representatives or the U.S. Senate, acting as science advisors, researchers, and staff consultants to members of Congress or Congressional committees. An annual stipend is paid by the sponsoring association. AVMF http://www.avmf.org AAAS http://www.aaas.org

There are 29 Sea Grant Colleges across the United States (associated with Land Grant Colleges) that offer Sea Grant Fellowships, where university scientists, educators, and outreach specialists are competitively chosen to work on Capitol Hill, on either House or Senate staff, in positions sponsored by Sea Grant, for as long as 1 year. Information on these fellowships can be accessed at: http://www.nsgo.seagrant.org


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Foundations

The Foundation Directory, for many years available at libraries, is now available online, and has listings by state and subject matter for private foundations offering grants to nonprofit organizations for special projects and operating expenses. Find the directory at: http://www.fconline.fdncenter.org Grants

The Grantsnet Web site is of great assistance in accessing grantors, as well as in providing tips on grant writing, career development, and foundation news. The site is accessed at: http://www.grantsnet.org

Federal Government Listings Federal jobs listing: Federal Office of Personnel Management http://www.usajobs.opm.gov

U.S. federal government listings: National Marine Fisheries Service, Silver Spring, MD http://www.nmfs.gov U.S. Agency for International Development, Washington, D.C. http://www.usaid.gov U.S. Department of Agriculture, Beltsville, MD http://www.usda.gov U.S. Department of the Interior, Washington, D.C. http://www.doi.gov U.S. Environmental Protection Agency, Washington, D.C. http://www.epa.gov U.S. Fish and Wildlife Service, Washington, D.C. http://www.fws.gov U.S. Geological Service (research arm of the Department of the Interior), Washington, D.C. http://www.usgs.gov National Park Service, Washington, D.C. http://www.nps.gov

Federal listings abroad: Canadian Department of Fisheries and Oceans http://www.ncr.dfo.ca

Miscellaneous Electronic Resources* A number of the organizations listed here also offer funds for research, as well as general veterinary and/or specific marine mammal medical information. Argus Clearinghouse http://www.clearinghouse.net/ * In alphabetical order; in the United States and abroad.

For subject-oriented topics, including science


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AVMA’s NOAH http://www.avma.org/network.html

Network of Animal Health

Cetacean Research Unit http://www.whalecenter.or

The Whale Center of New England

College of the Atlantic http://www.coa.edu/internships

Marine mammal courses and internships

Dalhousie University Whale Laboratory http://is.dal.ca/~whitelab/index.htm

Publications, information, and programs

Duke University Marine Mammal Laboratory http://www.env.duke.edu/marinelab/ marine.html

Marine resources, biomedical information, and library

Eckerd College Marine Mammal Courses http://www.eckerd.edu

Marine academic courses and programs

Institut Maurice Lamontagne http://www.qc.dfo-mpo.gc.ca/iml

Canadian oceans and fisheries information (French and English)

International Association for Bear Research http://www.bearbiology.com

Specific scientific information on bears (including polar bears)

International Biodiversity Measuring Course http://www.si.edu/simab/biomon.htm

Standardized protocols for biodiversity monitoring

International Marine Animal Trainers Association http://www.imata.org

Marine mammal science and public display

International Marine Mammal Association, Inc. http://www.imma.org

Marine mammal conservation and news

International Whaling Commission http://ourworld.compuserve.com/ homepages/iwcoffice

International convention for regulation of whaling

Ionian Dolphin Project http://www.tethys.org

Tethys Research Institute (Italian and English)

Manatee Awareness Coalition http://www.fmri.usf.edu/mammals.htm

Protecting Florida’s marine resources

Marine Mammal Careers (see also Chapter 7, Careers) http://www.seaworld.org/careers

SeaWorld

http://www.pegasus.cc.uct.edu/~smm

Society for Marine Mammalogy

http://www.rsmas.miami.edu/iof

International Oceanographic Foundation

Marine Mammal and Seabirds Course http://www.unb.ca/web/huntsman

University of New Brunswick, Canada

National Marine Educators Association http://www.marine-ed.org

Marine education, science, and research


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National Marine Mammal Laboratory http://nmml.afsc.noaa.gov

Marine mammal research in Northwest United States

North Atlantic Marine Mammal Commission http://www.nammco.no

Norway, Iceland, and Greenland marine mammal conservation and management

North Pacific Marine Mammal Research Consortium http://www.marinemammal.org

Bering Sea marine mammal research

Polar Bears Alive http://www.polarbearsalive.org

Polar bear and Arctic habitat information

Seal Conservation Society http://www.greenchannel.com/tec

Marine mammal welfare and conservation

Universita degli Studi di Pavia http://www.unipv.it/cibra

Marine mammal information (Italian and English)

Whales on the Net http://whales.magna.com.au/home.html

Cetacean information

Wildlife Disease Association http://www.vpp.vet.uga.edu/wda

Wildlife diseases, including marine mammals

Meetings and Proceedings on CD-ROM The following association annual meetings have aquatic animal medicine sessions, and proceedings of each meeting are available on CD-ROM. American Veterinary Medical Association (each year in July) Environmental Affairs, Aquatic Medicine, Public Health Sessions http://www.avma.org North American Veterinary Conference (each year in February in Orlando, FL) Aquatic Medicine, Wildlife Health Sessions http://vetshow.com/navc International Association for Aquatic Animal Medicine (each year in May) Aquatic Animal Medicine http://www.iaaam.org Western States Veterinary Conference (each year in February in Las Vegas, NV) Aquatic Medicine, Wildlife Health Sessions http://www.wvc.org

Electronic Addresses for Other Chapters in This Book Other pertinent Web sites specific to the scientific topics in each chapter of this book are noted in those chapters. You are directed to Chapter 7 (Careers) for information on continuing education opportunities in marine mammal medicine, as well as for a list of scientific societies and membership organizations related to marine mammal medicine. The Diagnostic Imaging Section of this book (Chapters 24 through 28) also contains a number of relevant technical Web site addresses.


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Disclaimer Because the number of Web sites related to marine mammal medicine is growing exponentially, the author cannot take responsibility for the complete exactness of the Internet addresses in this chapter. Although Web access to each site mentioned in this chapter was accomplished multiple times, be advised that Web site addresses change. To access the information if Web site addresses do change, we have provided the full organizational name and brief subject contents for each item in this chapter in order for you to conduct a search for the particular item of interest through standard search engines on the Web. The author, in accessing Internet Web sites in preparation of this chapter, has attempted to weed out those sites that are not of apparent high quality and/or value.

Conclusions One thing is certain, however. If you access the marine mammal medicine, conservation, and information sites included in this chapter, you will be better educated, not only in how to access the information, but also in how to read it with a critical eye and utilize it to your greatest advantage. The future of World Wide Web–based information systems is better designed Web sites, with consistency across veterinary medical information sites. In addition, the use of the Internet takes practice, just as any professional endeavor. The more you use the Web to access critical marine mammal resources and the more you attend seminars and continuing education sessions at conventions on accessing the Web, the better prepared you will be to manage and learn from the information you receive from the Internet. If we do this, along with our daily clinical practice and scientific reading, the marine mammals in our care will receive the best diagnostic and therapeutic approaches we can gather and implement.

References AAFP (American Academy of Family Practice), 1999, Computer Zoo, AAFP, Annual Meeting, Orlando, FL, 13 pp. BioMedicina, 1999, Medicine on the Internet: Surgery and ophthalmology in the information age, BioMedicina, 2(6): 295–298. Monti, D.J., 2000, SNOMED browser latest in informatics, J. Am. Vet. Med. Assoc. 216(7): 1049.


II Anatomy and Physiology of Marine Mammals



9 Gross and Microscopic Anatomy Sentiel A. Rommel and Linda J. Lowenstine

Introduction The California sea lion (Zalophus californianus) (Figure 1), Florida manatee (Trichechus manatus latirostris) (Figure 2), harbor seal (Phoca vitulina) (Figure 3), and bottlenose dolphin (Tursiops truncatus) (Figure 4) are used in this chapter to illustrate gross anatomy. These species were selected because of their availability and the knowledge base associated with them.* Gross anatomy of the sea otter (Enhydra lutra) is presented in Chapter 44 covering medical aspects of that species. Illustrations of the (A) external features, (B) superficial skeletal muscles, (C) relatively superficial viscera with skeletal landmarks, (D) circulation, body cavities, and some deeper viscera, and (E) skeleton are presented as five separate “layers” on the same page for each of the four species. These illustrations, based on dissections by one of the authors (S.A.R.), are of intact carcasses and thus help show the relative positions of organs in the live animals. The major lymph nodes are illustrated, but to simplify the illustrations, most are not labeled. The drawings represent size, shape, and position of organs in a healthy animal; the skeleton is accurately placed within the soft tissues and body outline. The scale of the drawings is the same for each species so that vertical lines can be used to compare features on all five; a photocopy onto a transparency would allow the reader to compare layers directly. Names of structures are labeled with three-letter abbreviations.** A brief figure legend helps the reader apply basic veterinary anatomical knowledge to the marine mammals illustrated. The style found in Miller’s Anatomy of the Dog (Evans, 1993) is followed as much as possible. Most technical terms follow the Illustrated Veterinary Anatomical Nomenclature by Schaller (1992). Recent comparative work on anatomy of marine mammals is found in Pabst et al. (1999), Rommel and Reynolds (2000; in press), and Reynolds et al. (in press). Older but valuable anatomical works include Murie (1872; 1874), Schulte (1916), Howell (1930), Fraser (1952), Slijper (1962), Green (1972), St. Pierre (1974), Bonde et al. (1983), King (1983), and Herbert (1987).

*A set of illustrations of a mysticete would be valuable, but as space is limited and they are less likely to be under veterinary care, we chose an odoctocete; the skeletal anatomy of the right whale (Eubalaena glacialis) is compared with that of other marine mammals in Rommel and Reynolds (in press). **Abbreviations in the text use capital letters to refer to the label on the structure. The first letter refers to the layer (A being external features at the top and E the skeleton) followed by a hyphen and then the abbreviation of the structure. For example, D-HAR refers to the heart on layer D.

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FIGURE 1 Left lateral illustrations of a healthy California sea lion (Zalophus californianus). Based on dissections by S.A.R., with details and nomenclatures from the literature: Murie, 1874; Howell, 1930; English, 1976a. Thanks to Rebecca Duerr for many helpful suggestions. (© Copyright S. A. Rommel. Used with permission of the illustrator.) (Layer A) External features. The following abbreviations are used as labels: ANG = angle of mouth; ANS = anus; AXL = axilla, flipperpit; CAL = calcaneus, palpable bony feature; EAR = external auditory opening, ear; EYE = eye; INS = cranial insertion of the extremity; flipper, fin, and/or fluke; NAR = naris; OLC = olecranon, palpable bony feature; PAT = patella, palpable bony feature; PEC = pectoral limb, fore flipper; PEL = pelvic limb, hind flipper; PIN = pinna, external ear (as opposed to external ear opening); SCA = dorsal border of the scapula, palpable (sometimes grossly visible) bony feature; TAI = tail; UMB = umbilicus; UNG = unguis, finger and toe nails; U/G = urogenital opening; VIB = vibrissae. (Layer B) The superficial skeletal muscles. The layer of skeletal muscles just deep to the blubber and panniculus muscles. The following abbreviations are used as labels: ANS = anus; BIF = femoral biceps; BRC = brachiocephalic; DEL = deltoid; DIG = digastric; EAM = external auditory meatus; EXT = external oblique; FAS = fascia; F,S,B&P = fur, skin, blubber, and panniculus muscle (where present) cut along midline; GLU = gluteals; LAT = latissimus dorsi; MAM = mammary gland; MAS = masseter; PECp = deep (profound) pectoral; PECs = superficial pectoral; REC = rectus abdominis; SAL = salivary gland; SER = serratus; nipple; STC = sternocephalic; TFL = tensor fascia lata; TMP = temporalis; TRAc = trapezius, cervical portion; TRAt = trapezius, thoracic portion; TRI = triceps brachii; UMB = umbilicus. (Layer C) The superficial internal structures with “anatomical landmarks.” This perspective focuses on relatively superficial internal structures; the other important bony or soft “landmarks” are not necessarily visible from a left lateral view, but they are useful for orientation. The relative size of the lung represents partial inflation—full inflation would extend the lung margins to the distal tips of ribs. The female is illustrated because there is greater variation in uterine anatomy than in testicular and penile anatomy; note, however, that only the sea lion (of the illustrated species) is scrotal (actually the sea lion testes migrate into the scrotum in response to environmental temperature). The following abbreviations are used as labels (structures in midline are in type, those off-midline are in italics): ANS = anus; AXL lnn = axillary lymph nodes; BLD = urinary bladder; F,S&B = fur, skin, blubber (cut at midline); HAR =heart; HYO = hyoid apparatus; INT = intestines; ILC = lliac crest; KID = left kidney; LIV = liver; LUN = lung (note that the lung extends under the scapula); MAN = manubrium of the sternum; OVR = left ovary; PAN = pancreas; PAT = patella; PSC ln = prescapular lymph nodes; RAD = radius; REC = rectum; SAL = salivary glands; SCA = scapula; SIG ln = superficial inguinal lymph node; SPL = spleen; STM = stomach; TIB = tibia; TMP = temporalis; TRA = trachea; TYR = thyroid gland; TYM = thymus gland; ULN = ulna; VAG = vagina. (Layer D) A view slightly to the left of the midsagittal plane illustrating the circulation, body cavities, and selected organs. Note that the diaphragm separates the heart and lungs from the liver and other abdominal organs. The following abbreviations are used as labels (structures on the midline are in normal type, those off-midline are in italics): AAR = aortic arch; ADR = adrenal gland; ANS = anus; AOR = aorta; ARH = aortic hiatus; AXL = axillary artery; BIF = tracheobronchial bifurcation; BLD = urinary bladder; BRC = bronchus; BRN = brain; CAF = caval foramen; CAR = carotid artery; caMESa = caudal mesenteric artery; CEL = celiac artery; CRZ = crus of the diaphragm; crMESa = cranial mesenteric artery; CVC = vena cava, between diaphragm and heart; DIA = diaphragm, cut at midline, extends from crura dorsally to sternum ventrally; ESO = esophagus (to the left of the midline cranially, on the midline caudally); ESH = esophageal hiatus; F,S&B = fur, skin, blubber (cut at midline); HAR = heart; HYO = hyoid bones; KID = right kidney; LIV = liver, cut at midline; LUN = right lung between heart and diaphragm; MAN = manubrium of sternum; OVR = left ovary; PAN = pancreas; PUB = pubic symphysis; PULa = pulmonary artery, cut at hilus of lung; PULv = pulmonary vein, cut at hilus of lung; REC = rectum; REN = renal artery; SPL = spleen; STM = stomach; STR = sternum, sternabrae; TNG = tongue; TRA = trachea; TYM = thymus gland; TYR = thyroid gland; UMB = umbilicus; UTR = uterus; VAG = vagina; VRT- vertebral artery; XIP = xyphoid process of the sternum. (Layer E) The skeleton. Regions of the vertebral column (cervical, thoracic, lumbar, sacral, and caudal) are abbreviated (in lower case) as cer, tho, lum, sac, and cau, respectively, and are used as modifiers after an abbreviation in caps and a comma. If a specific vertebra is labeled, it will be represented by a capitalized first letter (for caudal, Ca will be used) and the vertebral number, i.e., first cervical = C1. The following abbreviations are used as labels: CAL = calcaneus; CAN = canine tooth (not present in cetaceans or manatees); DIG = digits; FEM = femur; FIB = fibula; HUM = humerus; HYO = hyoid bones; ILC = iliac crest of the pelvis; LRB = last, or caudalmost, rib; MAN = mandible; MNB = manubrium, the cranialmost bony part of the sternum; NSP = neural spine (spinous process), e.g., thoracic neural spines = NSP, tho; OLC = olecranon; ORB = orbit; PAT = patella; RAD = radius; SCA = scapula; STN = sternum, composed of individual sternabrae; SRB = sternal ribs, costal cartilages; TIB = tibia; TMF = temporal fossa; TPR = transverse process, e.g., TPR, C1 = transverse process of the first cervical vertebra; ULN = ulna; VBR = vertebral ribs; ZYG = zygomatic arch.


131

Gross and Microscopic Anatomy

OLC

PIN SCA

EAR

PAT

EYE ANS CAL TAI

UNG

NAR ANG

VIB

INS

A

U/G PEL INS

U/G

UMB

AXL UNG PEC LAT

TRAt EAM

BRC

SER

FAS

TRAc

TMP

F, S & B TFL GLU BIF ANS

MAS

DIG

SAL STC

B

F, S, B & P MAM

DEL

EXT

PECs

PECp TRI

LUN

AXL Inn 1-3

HYO

TMP

UMB

REC

F, S & B

SCA

PSC Inn

PAN

KID ILC REC

EYE

ANS

VAG

SAL TYR

C

SIGIn

TRA MAN

HUM

HAR

TYM

LIV

SPL

STM

OVR

INT

PAT BLD

TIB

RAD ULN c Rommel 2000

ESO

ESO AAR

F, S & B

BRN

CAF CVC DIA CEL crMESa AOR ESH ESO LUN ARH BRC PAN CRZ

ADR REN

KID

caMESa PUB

TNG

ANS

HYO

VAG

TYR

D

UTR

CAR TRA

BLD

VRT BIF MAN

TMF

OVR

REC

AXL TYM PULa PULv SPL HAR STR DIA XIP LIV STM UMB NSP tho

NSP, cer

VBR

LRB

SCA

NSP, Ium ILC

ORB

NSP, cau

CAL CAN MAN

E

ZYG

HYO TPR, C1

TIB PAT

MNB

FIB

FEM DIG

HUM

SRB

OLC STN RAD ULN DIG


132


FIGURE 2 Left lateral illustrations of a healthy Florida manatee (Trichechus manatus latirostris). Based on dissections by S.A.R., with details and nomenclatures from the literature: Murie, 1872; Domning, 1977; 1978; Rommel and Reynolds, 2000. Thanks to D. Domning for suggestions on the muscle illustration. (© S. A. Rommel. Used with permission of the illustrator.) (Layer A) External features. The following abbreviations are used as labels: ANG = angle of mouth; ANS = anus; AXL = axilla; EAR = external auditory opening, ear; EYE = eye; FLK = fluke entire caudal extremity in manatees; flukes = entire caudal extremity in dugongs; INS = cranial insertion of the extremity, flipper and/or fluke; NAR = naris; OLC = olecranon, palpable bony feature; PEC = pectoral limb, flipper; PED = peduncle, base of tail, between anus and fluke; SCA = dorsal border of the scapula, palpable bony feature in emaciated individuals; UMB = umbilicus; UNG = unguis, fingernails; U/G = urogenital opening; VIB = vibrissae. (Layer B) The superficial skeletal muscles. The layer of skeletal muscles just deep to the blubber and panniculus muscles. The following abbreviations are used as labels: ANS = anus; CEP = cephalohumeralis; DEL = deltoid; EXT = external oblique; FAS = fascia; S,B&P = skin, blubber, and panniculus muscle (where present) cut along midline; IIN = internal intercostals; ILC = iliocostalis; ITT = intertransversarius; LAT = latissimus dorsi; LEN = levator nasolabialis; LON = longissimus; MAM = mammary gland, in axillary region, thus partly hidden under the flipper; MEN = mentalis; MND = mandibularis; PAN = panniculus, illustrated using dotted lines, is a robust and dominant superficial muscle; a layer of blubber is found on both the medial and lateral aspects of this muscle; REC = rectus abdominis; SLT = mammary slit, nipple; SPC = sphincter colli; SVL = sarcoccygeus ventralis lateralis; TER = teres major; TMP = temporalis; TRA = trapezius; TRI = triceps brachii; UMB = umbilicus, XIN = external intercostals. (Layer C) The superficial internal structures with “anatomical landmarks.” This perspective focuses on relatively superficial internal structures. Skeletal elements are included for reference, but not all are labeled. The left kidney (not visible from this vantage in the manatee) is illustrated. The relative size of the lung represents partial inflation. The following abbreviations are used as labels: ANS = anus; BLD = urinary bladder (dotted, not really visible in this view); BVB = brachial vascular bundle; CHV = chevrons, chevron bones; EYE = the eye (note how small it is); HAR = heart; HUM = humerus; INT = intestines; note the large diameter of the large intestines; KID = left kidney, not visible from this vantage in the manatee; LIV = liver; LUN = lung (note lung extends under scapula, and over heart); OVR = left ovary; PEL = pelvic vestige; RAD = radius; SAL = salivary gland; S&B = skin and blubber; SCA = scapula; SIG ln = superficial inguinal lymph node; S,B&P = skin, blubber, and panniculus muscle, cut at midline; STM = stomach; TMJ = temporomandibular joint; TYM = thymus gland; ULN = ulna; UMB = umbilical scar; UTR = uterine horn; VAG = vagina. (Layer D) A view slightly to the left of the midsagittal plane illustrates the circulation, body cavities, and selected organs. Note that the diaphragm of the manatee is unique and that the distribution of organs and the separation of thoracic structures from abdominal structures requires special consideration. The following abbreviations are used as labels (structures on the midline are in normal type, those off-midline are in italics): AAR = aortic arch; ADR = left adrenal gland; ANS = anus; AOR = aorta; AXL = axillary artery; BLD = urinary bladder; BRN = brain; BVB = brachial vascular bundle (cut); CAF = caval foramen; CAR = carotid artery; CDG = cardiac gland; CEL = celiac artery; CER = cervix; CHV = chevron bones; CRG = cardiac gland; CVB = caudal vascular bundle; DUO = duodenum; ESO = esophagus (to the left of the midline cranially, on the midline caudally); EXI = external iliac artery; HAR = heart; KID = right kidney; LIV = liver, cut at midline; OVR = right ovary; PAN = pancreas; PULa = pulmonary artery, cut at hilus of lung; PULv = pulmonary vein, cut at hilus of lung; REC = rectum; REN = renal artery; S&B = skin and blubber; SKM = skeletal muscle; SM&B = skin, muscle, and blubber (cut at midline); SPL = spleen; STM = stomach; STR = sternum; TNG = tongue; TRA = trachea; TRS = transverse septum; TYM = thymus gland; TYR = thyroid gland; UMB = umbilical scar; UTR = uterus; VAG = vagina. (Layer E) The skeleton. Regions of the vertebral column (cervical, thoracic, lumbar, sacral, and caudal), are abbreviated (in lowercase) as cer, tho, lum, sac, and cau, respectively, and are used as modifiers after an abbreviation in caps and a comma. If a specific vertebra is labeled, it will be represented by a capitalized first letter (for caudal, Ca will be used) and the vertebral number, i.e., first cervical = C1. The following abbreviations are used as labels: CHV = chevrons, chevron bones; DIG = digits, columns of finger bones; HUM = humerus; HYO = hyoid apparatus; HYP = hypapophysis, ventral midline vertebral process; LRB = last, or caudalmost, rib; LVR = last, or caudalmost, vertebra; MAN = mandible; NSP = neural spine (spinous process), e.g., thoracic neural spines = NSP, tho; OLC = olecranon; ORB = orbit; PEL = pelvic bone; RAD = radius; SCA = scapula; STN = sternum, if sternabrae are commonly fused; SBR = sternal ribs, costal cartilages; TMF = temporal fossa; TPR = transverse process, C1; ULN = ulna; VBR = vertebral ribs; XNR = external (bony) nares; XIP = xyphoid process, cartilaginous caudal extension of the sternum; ZYG = zygomatic process of the squamosal.


133


OLC SCA FLK

EAR PED EYE NAR

INS

VIB

A

ANS

INS

ANG

AXL

U/G UMB

PEC

UNG

U/G

ILC

XIN

IIN

LAT

LON

TER

S, B, & P

TRA

CEP

ITT FAS

TEM LEN

SVL MEN

SPC DEL

MND

ANS

TRI MAM

SLT

B

S, B, & P

UMB

REC

S&B

LUN

KID (not visible)

LUN LUN

SCA

PAN

EXT

UTR

LIV

OVR PEL

TMJ

S&B

SAL EYE

CHV HUM

ANS

TYM

C

BVB

STM CRG HAR INT (lg)

RAD

INT (sml) UMB

SIG In

BLD INT (lg) S, B & P

ULN

PULa AXL AAR TRA

PULv

ESO AOR

CRG

AOR

CEL

ADR

c Rommel 2000

SKM

REN

OVR

EXI CVB

TYR S&B

BRN CAR

CHV BVB

TNG

SKM TYM

D

ANS REC

HAR STR CAF TRS LIV

STM

SPL DUO PAN UMB

SM&B

KID

UTR

BLD

VAG

CER

NSP, tho SCA TPR, C1

NSP, lum

NSP, cer

LVR

TPR, Ca1

HYO

NSP, ca

TMF ZYG XNR

ORB CHV MAN

HUM STN

E

PEL OLC

RAD

DIG

ULN

SBR

LRB HYP

VBR


134


FIGURE 3 Left lateral illustrations of a healthy harbor seal (Phoca vitulina). Based on dissections by S.A.R., with details and nomenclatures from the literature: Howell, 1930; Huber, 1934; Bryden, 1971; Tedman and Bryden, 1981; Rommel et al., 1998; Pabst et al., 1999. (© Copyright S. A. Rommel. Used with permission of the illustrator.) (Layer A) External features. The following abbreviations are used as labels: ANG = angle of mouth; ANS = anus; AXL = axilla; CAL = calcaneus, palpable bony feature; EAR = external auditory opening, ear; EYE = eye; INS = cranial insertion of the flipper; NAR = naris; OLC = olecranon, palpable bony feature; PAT = patella, palpable bony feature; PEC = pectoral limb, fore flipper; PEL = pelvic limb, hind flipper; SCA = dorsal border of the scapula, palpable bony feature; TAI = tail; UMB = umbilicus; UNG = unguis, finger and toe nails; U/G = urogenital opening; VIB = vibrissae. (Layer B) The superficial skeletal muscles. The layer of skeletal muscles just deep to the blubber and panniculus muscles. The following abbreviations are used as labels: ANS = anus; BIF = femoral biceps; BRC = brachiocephalic; DEL = deltoid; DIG = digastric; EAM = external auditory meatus; EXT = external oblique; FAS = fascia; F,S&B = fur, skin, blubber, and panniculus muscle (where present) cut along midline; GLU = gluteals; GRA = gracilis; LAT = latissimus dorsi; MAM = mammary gland; MAS = masseter; PAR lnn = parotid lymph nodes (ln for a single lymph node); PECa = ascending pectoral, extends over the patella and part of hind limb; PECs = superficial, pectoral; PECp = deep (profound) pectoral; REC = rectus abdominis; SAL = salivary gland; SEM = semitendinosus; SER = serratus; STC = sternocephalic; STH = sternohyoid; TFL = tensor fascia lata; TMP = temporalis; TRAc = trapezius, cervical portion; TRAt = trapezius, thoracic portion; TRI = triceps brachii; UMB = umbilicus. (Layer C) The superficial internal structures with “anatomical landmarks.” A view focused on relatively superficial internal structures visible from that perspective; the other important bony or soft “landmarks” are not necessarily visible from a left lateral view, but they are useful for orientation. The relative size of the lung represents partial inflation—full inflation would extend margins to distal tips of ribs. The following abbreviations are used as labels: ANS = anus; AXL = axillary lymph node; BLD = urinary bladder; EYE = eye; FEM = femur; FIB = fibula; HAR = heart; HUM = humerus; HYO = hyoid apparatus; INT = intestines; ILC = lliac crest; KID = left kidney; LIV = liver; LUN = lung; MAN = manubrium of the sternum; OLE = olecranon; OVR = left ovary; PAN = pancreas; PAT = patella; PRE = presternum, cranial sternal cartilage; PSC ln = prescapular lymph node; RAD = radius; REC = rectum; SAL = salivary glands; SIG ln = superficial inguinal lymph node; SCA = scapula; SPL = spleen; STM = stomach; TMJ = temporomandibular joint; TIB = tibia; TRA = trachea; TYR = thyroid gland; TYM = thymus gland; ULN = ulna; UMB = umbilical scar; UTR = left uterine horn; VAG = vagina; XIP = xiphoid. (Layer D) A view slightly to the left of the midsagittal plane illustrates the circulation, body cavities, and selected organs. Note that the diaphragm separates the heart and lungs from the liver and other abdominal organs. The following abbreviations are used as labels (structures on the midline are in normal type, those off-midline are in italics): AAR = aortic arch; ADR = left adrenal gland; ANS = anus; AOR = aorta; AXL = axillary artery; BCT = left brachiocephalic trunk; BRC = left bronchus as it enters the lung; BLD = urinary bladder; BRN = brain; CAF = caval foramen, with caval sphincter; CAR = carotid artery; CEL = celiac artery; CER = cervix; CVC = caudal vena cava; CRZ = left crus of the diaphragm; DIA = diaphragm, cut at midline, extends from crura dorsally to sternum ventrally; ESO = esophagus (to the left of the midline cranially, on the midline caudally); ESH = esophageal hiatus; EXI = external iliac artery; F,S&B = fur, skin, and blubber, plus panniculus where appropriate, cut on midline; HAR = heart; HPS = hepatic sinus within liver; KID = right kidney; LIV = liver, cut at midline; LUN = lung, right lung between heart and diaphragm; MAN = manubrium of sternum; caMESa = caudal mesenteric artery; crMESa = cranial mesenteric artery; OVR = ovary; PAN = pancreas; PUB = pubic symphysis; PULa = pulmonary artery, cut at hilus of lung; PULvv = pulmonary veins, cut at hilus of lung; REC = rectum; REN = renal artery; SKM = skeletal muscle; SPL = spleen; STM = stomach; STR = sternum made up of individual sternabrae; TNG = tongue; TRA = trachea; TYM = thymus gland; TYR = thyroid gland; UMB = umbilicus; UTR = uterus; VAG = vagina; XIP = xyphoid process of the sternum. (Layer E) The skeleton. Regions of the vertebral column (cervical, thoracic, lumbar, sacral, and caudal) are abbreviated (in lower case) as cer, tho, lum, sac, and cau, respectively, and are used as modifiers after an abbreviation in caps and a comma. If a specific vertebra is labeled, it will be represented by a capitalized first letter (for caudal, Ca will be used) and the vertebral number, i.e., first cervical = C1. The following abbreviations are used as labels: CAL = calcaneus; CAN = canine tooth; DIG = digits; FEM = femur; FIB = fibula; HUM = humerus; HYO = hyoid bones; ILC = iliac crest of the pelvis; LRB = last, or caudalmost, rib; LVR = last, or caudalmost, vertebra; MAN = mandible; MNB = manubrium, the cranialmost bony part of the sternum; NSP = neural spine (spinous process), e.g., thoracic neural spines = NSP, tho; OLC = olecranon; ORB = orbit; PAT = patella; PRS = presternum, cartilaginous extension of the sternum, particularly elongate in seals; PUB = pubic symphysis; RAD = radius; SCA = scapula; SBR = sternal ribs, costal cartilages; TIB = tibia; TMF = temporal fossa; TPR = transverse process, e.g., TPR, C1 = transverse process of the first cervical vertebra; ULN = ulna; VBR = vertebral ribs; XNR = external (bony) nares, nasal aperture of the skull; XIP = xyphoid process, cartilaginous caudal extension of the sternum; ZYG = zygomatic arch.


135


AXL OLC

EAR SCA EYE

CAL TAI

ANS

U/G

NAR

VIB ANG

A

INS

PEL UNG

PAT U/G INS

UNG

PEC

UMB FAS

LAT

F, S & B TFL

TRI

EAM

TMP

GLU

BRC TRAt

TRAc

SAL

BIF SEM

ANS

MAS

DIG PAR Inn

B

STH GRA

STC F, S & B DEL PECs SER

PECa

UMB

F, S & B

REC

KID

LUN HYO

TMJ

OVR

OLE

SAL

ILC

SCA

PSC In

EYE

EXT

MAM PECp

FEM FIB REC

ANS

TYR

C

U/G

TRA TIB PRE

SIN In

MAN

BLD

HUM TYM

PAT AXL In

RAD

ULN

XIP

LIV

SPL

STM INT

UMB PAN

UTR

HAR c Rommel 2000

CAR

BRN

ESO

SKM VRT

AAR

PULa ESO

BRC

LUN

ESH DIA CEL crMESa CAF AOR

ADR

CRZ

KID

REN caMESa

EXI F, S & B REC

ANS VAG

TNG TYR

D

CER PUB

TRA MAN

BLD

AXL

TYM

BCT

STR

PULvv HAR CVC DIA XIP HPS

LIV

STM VBR

NSP, tho

NSP, C2

ORB

OVR

F, S & B

UTR

LRB NSP, lum

OLC TMF

SPL UMB

PAN

ILC

SCA

NSP, cau CAL

XNR

LVR

CAN MAN ZYG

E

HYO TPR,C1

PUB FIB

PRS TIB

MNB PAT

HUM RAD

FEM ULN

XIP DIG

SBR

DIG

0839_frame_C09 Page 136 Wednesday, May 23, 2001 10:42 AM

136


FIGURE 4 Left lateral illustrations of a healthy bottlenose dolphin (Tursiops truncatus). Based on dissections by S.A.R. with details and nomenclatures from the literature: Howell, 1930; Huber, 1934; Fraser, 1952; Slijper, 1962; Mead, 1975; Strickler, 1978; Klima et al., 1980; Pabst, 1990; Rommel et al., 1998; Pabst et al., 1999. Thanks to T. Yamada for suggestions on the muscle illustration. (© S. A. Rommel. Used with permission of the illustrator.) (Layer A) External features. The following abbreviations are used as labels: ANG = angle of mouth; ANS = anus; AXL = axilla; BLO = blowhole, external naris in dolphin; EAR = external auditory opening, ear; EYE = eye; FIN = dorsal fin; FLK = flukes (entire caudal extremity in cetaceans); INS = cranial insertion of the extremity; flipper, fin, and/or fluke; NOC = fluke notch in dugongs and in most cetaceans; PEC = pectoral limb, flipper; PED = peduncle, base of tail, between anus and flukes; MEL = melon; SCA = dorsal border of the scapula, palpable bony feature in emaciated dolphins; SNO = snout, cranial tip of upper jaw; UMB = umbilicus; U/G = urogenital opening. (Layer B) The superficial skeletal muscles. The layer of skeletal muscles just deep to the blubber and panniculus muscles. Note that the large muscles ventral to the dorsal fin are surrounded by a tough connective tissue sheath (Pabst, 1990). The following abbreviations are used as labels: ANS = anus; BLO = blowhole; DEL = deltoid; DIG = digastric; EAM = external auditory meatus; EPX = epaxial muscles, upstroke muscles; EXT = external oblique; HYP = hypaxialis; HPX = hypaxial muscles, downstroke muscles; ILI = iliocostalis; INT = internal oblique; ISC = oschium; ITTd = intertransversarius caudae dorsalis; ITTv = intertransversarius caudae ventralis; LAT = latissimus dorsi; LEV = levator ani; LON = longissimus; MAM = mammary gland; MAS = masseter; MUL = multifidus; PECp = deep (profound) pectoral; PSC ln = presacpular lymph node; REC = rectus abdominis; RHO = rhomboid; ROS = rostral muscles; S,B,&P = skin, blubber, and panniculus muscle (where present) cut along midline; SER = serratus; SLT = mammary slit, nipple; SPL = splenius; STE = sternohyoid; STM = sternomastoid; TER = teres major; TMP = temporalis; TRAd = trapezius dorsalis; TRAc = trapezius cranialis; TRI = triceps brachii; UMB = umbilicus. (Layer C) The superficial internal structures with “anatomical landmarks.” The relative size of the lung represents partial inflation—full inflation would extend margins to distal tips of ribs. The following abbreviations are used as labels: ANS = anus; BLD = urinary bladder; BLO = blowhole; EYE = eye; HAR = heart; HPX = hypaxial muscles; HUM = humerus; HYO = hyoid apparatus; INT = intestines; KID = left kidney; LIV = liver; LUN = lung (note that it extends beneath the scapula); MEL = melon; OVR = left ovary; PEL = pelvic vestige; PSC ln = prescapular lymph node; PUL ln = pulmonary lymph node, unique to cetaceans; RAD = radius; REC = rectum; ROS = rostral muscles, to manipulate the melon; SAC = lateral diverticulae, air sacs in dolphin; S&B = skin and blubber; SCA = scapula; SKM = skeletal muscle; SPL = spleen; STM = stomachs; TMJ = temporomandibular joint; TRA = trachea; TYR = thyroid gland; ULN = ulna; UMB = umbilical scar; UOP = uterovarian plexus; URE = ureter; UTR = uterine horn; VAG = vagina. (Layer D) A view slightly to the left of the midsagittal plane illustrates the circulation, body cavities, and selected organs. Note that the diaphragm separates the heart and lungs from the liver and other abdominal organs. The following abbreviations are used as labels (structures on the midline are in normal type, those off-midline are in italics): AAR = aortic arch; ADR = left adrenal gland; ANS = anus; AOR = aorta; AXL = axillary artery; BLD = urinary bladder; BLO = blowhole; BRC = bronchus; BRN = brain; CAR = carotid artery; CEL = celiac artery; CER = cervix; CRZ = left crus of the diaphragm; CVB = caudal vascular bundle; DIA = diaphragm, cut at midline, extends from crura dorsally to sternum ventrally; ESO = esophagus (to the left of the midline cranially, on the midline caudally); ESH = esophageal hiatus; EXI = external iliac artery; FINaa = arteries arrayed along the midline of the dorsal fin; FLKaa = arterial plexus on dorsal and ventral aspects of each fluke; HAR = heart; KID = right kidney; LAR = larynx or goosebeak; LIV = liver, cut at midline; MEL = melon; OVR = right ovary; PAN = pancreas (hidden behind first stomach); PMX = premaxillary sac; PULa = pulmonary artery, cut at hilus of lung; PULv = pulmonary vein, cut at hilus of lung; REC = rectum; REN = renal artery; S&B = skin and blubber, panniculus where appropriate cut at midline; SKM = skeletal muscle; SPL = spleen; STM1 = forestomach; STM2 = main stomach; STM3 = pyloric stomach; STR = sternum, sternabrae; TNG = tongue; TRA = trachea; TYM = thymus gland; TYR = thyroid gland; UMB = umbilicus; UOP = right uterovarian vascular plexus in dolphin; URE = right ureter; UTR = uterus; VAG = vagina. (Layer E) The skeleton. Regions of the vertebral column (cervical, thoracic, lumbar, sacral, and caudal), are abbreviated (in lower case) as cer, tho, lum, sac, and cau, respectively, and are used as modifiers after an abbreviation in caps and a comma. If a specific vertebra is labeled, it will be represented by a capitalized first letter (for caudal, Ca will be used) and the vertebral number, i.e., first cervical = C1. The following abbreviations are used as labels: CHV = chevrons, chevron bones; DIG = digits; HUM = humerus; HYO = hyoid apparatus; LRB = last, or caudalmost, rib; LVR = last, or caudalmost, vertebra; MAN = mandible; NSP = neural spine; e.g., thoracic neural spines = NSP, tho; OLC = olecranon; ORB = orbit; PEL = pelvic vestige; RAD = radius; SCA = scapula; STR = sternum; SBR = sternal ribs, costal ribs; TMF = temporal fossa; ULN = ulna; VBR = vertebral ribs; XNR = external (bony) nares, nasal aperture of the skull; ZYG = zygomatic arch.


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INS

SCA

FIN

EAR BLO EYE PED MEL

FLK

SNO

ANG

INS

INS

A

ANS

U/G PEC UMB

AXL

PSC In SPL TRAc SEM

TRAd

LAT

RHO

MUL

NOC

U/G

LON ILI EPX

S&B

EAM BLO

MUL

LON

TEM

ITTd

ROS

MAS

DIG STE STM MAS DEL

B

PSC In SAC EYE

TRI PECp INF TER

SCA

LUN

LUN

SER REC

INT UMB

EXT

MAM

SLT

ANS

ITTv

ISC HYP HPX

S, B & P

SPL KID URE

OVR

BLO

REC S&B

MEL

SKM

ROS

TMJ

HYO

TRA TYR HUM PEL VAG ANS

RAD ULN

C

LIV PUL In

HAR

STM UMB

UTR INT HPX

S&B

SKM

BLD

UOP

REN

CAR TRA BRN PMX

ESO

BRC AAR PULa

CRZ PAN (hidden) CEL PULv ESH SKM SPL

FINaa

OVR

UOP

c Rommel 2000 AOR

BLO

EXI SKM

MEL

REC

S&B

CVB SKM

TNG

LAR TYR TYM

AXL

STR HAR

D

DIA LIV

STM 2 STM 3 STM 1

CER UMB

UTR ADR KID

URE

VAG ANS SKM S&B

FLKaa

BLD

NSP, tho

SCA NSP, C1&2

NSP, lum

TMF XNR

NSP, cau

ORB

MAN

LVR

ZYG

HYO

HUM PEL

RAD

E

OLC

STR ULN DIG

SBR

VBR

LRB

CHV


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Included is a section on microanatomy to introduce the microanatomical peculiarities of marine mammals to pathologists and thus aid them in performing routine histopathological examination of marine mammal tissues. The microscopic appearance of organs and tissues is presented following the gross anatomical descriptions. This information has been gathered from the examination of tissues submitted to the University of California Veterinary Medical Teaching Hospital Pathology Service over the last 20 years. These tissues were acquired from stranded marine mammals, such as California sea lions, harbor seals, northern elephant seals (Mirounga angustirostris), southern sea otters (Enhydra lutris nereis), and a few small odontocetes and gray whales (Eschrichtius robustus). Anatomical observations from the literature are also included and referenced. Previous reviews of microanatomy include Simpson and Gardner (1972), Britt and Howard (1983), and Lowenstine and Osborne (1990). Histological recognition of organs and tissues from marine mammals poses little problem for individuals acquainted with the microanatomy of terrestrial mammals. The patterns of degenerative, inflammatory, and proliferative changes observed in marine mammal tissues are also similar to those observed in domestic mammalian species. Knowledge of specific microanatomy is necessary, however, for subtle changes to be recognized.

External Features Consider the morphological features of the selected marine mammals. Streamlining and thermoregulation have caused changes in the appearance of marine mammals; these adaptations include the modification of appendages and other extremities for swimming, an increase in blubber for insulation, the development of axial locomotion, and the development of ascrotal testes (Pabst et al., 1999).

Sea Lions The otariids (fur seals and sea lions), represented by the California sea lion, are also called eared seals because they have distinct pinnae (A-PIN) associated with their external ear openings (A-EAR). Like other pinnipeds, sea lions have robust vibrissae (A-VIB) on their snouts. Fur and/or blubber help streamline and insulate their bodies. Otariids (and walruses) can assume distinctly different postures on land by rotating their pelves to position their pelvic (or hind) flippers (A-PEL) under their bodies. Note the presence of nails (unguis; A-UNG) on the extremities. Eared seals propel themselves with their pectoral (or fore) flippers (A-PEC) when swimming. The adult males of the sexually dimorphic California sea lion (and most other otariids) are much larger than the females. The teeth of sea lions are often stained dark brown or black in the absence of significant dental calculus. As in other carnivora, the nasal turbinates are well developed (Mills and Christmas, 1990).

Manatees The sirenians are represented by the Florida manatee. They lack hind limbs and have a dorsoventrally flattened fluke (A-FLK; note that it is flukes in cetaceans and dugongs and fluke in manatees). There is no dorsal fin, and the pectoral limbs or flippers are much more mobile than those of cetaceans—it is common to see manatees with their flippers folded across their chests or manipulating food into the mouth. The skin is rough and relatively thick and massive when compared with that of terrestrial mammals of the same body size. The skin is denser than water and contributes significantly to negative buoyancy (Nill et al., 2000). The vibrissae are robust but short (from wear), and the body hairs are fine but sparse, and give a nude



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appearance to the skin of the manatee. Although body hairs are sparse, they are uniquely innervated and might provide vibrational and other tactile sensations (Reep et al., 1999). The eyes (A-EYE) of manatees are small and, unlike the eyes of other mammals, close using a sphincter rather than distinct upper and lower eyelids.

Seals The phocids, or earless seals (also called hair seals), are represented by the harbor seal. They have vibrissae similar to those of a dog. Their nares (A-NAR) are located at the dorsal aspects of their snouts. Phocid eyes are typically large (C-EYE) when compared with those of other marine mammals. Note that the appearance of phocids is generally the same, whether they are in the water or on land. Phocids commonly tuck their heads back against the thoraxes, making the neck look shorter than it really is, and they locomote in the water by lateral undulation of their pelvic flippers (A-PEL). Their flippers have long curved nails (A-UNG). Some phocids have multiple cusps on the caudal teeth, which in some species are quite complex and ornate.

Dolphins The odontocetes are represented by the bottlenose dolphin. The cetaceans are characterized by the absence of pelvic limbs but are graced with large caudal structures called flukes (A-FLK). The melon (A-MEL) is a rostral fat pad that, together with elongated premaxillae and maxillae, gives the dolphin its “bottlenose.” The external nares are joined as a single respiratory opening at the blowhole (A-BLO), located at or near the apex of the skull. The externally smooth skin of dolphins has a thickened dermis, referred to as blubber. Some cetaceans also have dorsal fins (A-FIN), which are midline, nonmuscular, fleshy structures that may help stabilize them hydrodynamically. The keel of the peduncle (A-PED) provides streamlining and acts as a mechanical spring (Pabst et al., 1999). Cetaceans also have a pair of pectoral flippers that help them steer. Dolphins have facial hairs in utero but lose them at or near the time of birth (Brecht et al. 1997). Drawings contrasting features of the head and teeth of a representative porpoise and a representative dolphin appear in Reynolds et al. (1999). The unusual head of the sperm whale (Physeter macrocephalus) is described in detail by Cranford (1999). Dolphins have conical, pointed (when young and unworn) teeth. In contrast to dolphins, porpoises have flattened spade-shaped teeth and the lower, cranial margin of the melon extends all the way to the margin of the upper jaw or beak—there is no “bottle-shaped nose.” As dolphins age, their teeth wear down, as they are abraded by ingested material and each other; the name truncatus is derived from the truncated appearance of the teeth in the original specimen. The tongues of the bottlenose dolphin and some other odontocetes have elaborate cranial and lateral marginal papillae, which are important for nursing (Donaldson, 1977).

Microanatomy of the Integument The cetacean integument differs significantly from that of terrestrial mammals in that there are no hair follicles (save for a few on the snouts of some species) and no sebaceous or apocrine glands (Greenwood et al., 1974; Ling, 1974). The thick epidermis is nonkeratinizing, lacks a granular layer, and is composed primarily of stratum spinosum (stratum intermedium) with deep rete pegs. The basal layer has continuous mitoses. Continuous desquamation caused by water friction may account for the absence of a keratinized stratum corneum and the continuous cell replication in the basal layer. The papillary dermis is extremely well vascularized (Elsner et al., 1974). The reticular dermis grades into the fat-filled panniculus adiposus, creating a fatty


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layer referred to as the blubber layer. The blubber contains many collagen (fibrous) bundles and elastic fibers, and adipocytes are interspersed so that blubber thickness may not diminish significantly during catabolism of fat. The blubber layer is connected to the underlying musculature by loose connective tissue (subcutis). Pinnipeds, sea otters, and sirenians are haired (although hair density varies enormously from sea otters to walruses and sirenians), and therefore their skin is more similar to domestic mammals than is cetacean skin. The epidermis of these species is partially or entirely keratinizing. The stratum corneum is thickest on weight-bearing surfaces, such as the relatively glabrous ventral surfaces of fore and hind flippers, where the entire epidermis is quite thick. A stratum granulosum is present in phocids. Compound hair follicles consisting of a single guard hair follicle and several intermediate and underfur follicles are common, especially in fur seals and sea otters. Elephant seals, monk seals, and walruses, which lack underfur, all have simple hair follicles consisting of a single guard hair. Like terrestrial mammals, hair follicles of sea otters and pinnipeds are associated with well-developed sebaceous and apocrine (sweat) glands. Apocrine sweat glands are relatively large in the otariid seals, whereas the sebaceous glands are more prominent in the phocids. In densely haired regions of fur seals, the sweat glands enter the hair follicle above (distal) the sebaceous gland duct, but in sparsely haired species (such as the harp seal) and in sparsely haired areas of densely haired species, the pattern is reversed (Ling, 1974). Concentrations of glands vary with location on the animal, and patterns of gland distribution have not been fully described for all species. In some pinniped species, apocrine gland secretion may be more evolved for scent and olfactory communication than for thermoregulation (Greenwood et al., 1974). Hair follicles in all species are said to lack arrector pili muscles and have a fairly fixed angle relative to the skin surface. Vibrissae may be selectively heated by changes in blood flow (Mauck et al., 2000). The blubber layer is relatively thin in fur seals and sea otters; in these species, the pelage is presumed to provide primary insulation. The connective tissue in the pinniped dermis contains many elastic fibers. The reticular layer is thicker than the papillary layer. The lower portions of hair follicles extend into the deep reticular dermis and are often surrounded by adipose tissue in those species with a thick blubber layer. An interesting physiological phenomenon involving the marine mammal integument is the catastrophic cyclic molting that occurs in some phocids (Ling, 1974). Domestic mammals also tend to shed hair cyclically, but the stratum corneum is desquamated continuously, accompanied by continuous proliferation of the basal cell layer. In some phocids, basilar mitosis is seasonal, and the lipid-rich stratum corneum is parakeratotic and persists as a protective, presumably waterproof, sheet from one molt to the next. Prior to molt, a granular cell layer develops, and during molt, the surface epithelium is shed in great sheets along with the hair. In harp seals, this process is manifest grossly as small circular lesions that open and become confluent, leading to a drying-out and sloughing of the entire epidermal surface. Catastrophic molt has been best described histologically in the southern elephant seal (M. leonina) and is also evident in the northern elephant seal. Cyclic shedding or molt has also been seen in otariids but occurs more slowly, with shedding of the hair over several weeks or months. Mammary glands (B-MAM) are ventral, medial, and relatively caudal in most marine mammals, but they are axillary in sirenians. Cetaceans and some phocids have a single pair of nipples (B-SLT), but otariids and polar bears have two pairs of nipples. In cetaceans, the nipples are within mammary slits located lateral to the urogenital opening (note that some male cetaceans have distinct mammary slits). Detailed anatomy of the phocid mammary gland is described by Bryden and Tedman (1974) and Tedman and Bryden (1981).



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The Superficial Skeletal Muscles The skeletal muscles that are encountered when the skin, blubber,* and panniculus muscles are removed are illustrated in layer B of each figure. Note that the panniculus (B-PAN) is represented as dotted lines in the manatee because it is such a robust muscle, bordered on its lateral and medial aspects by “blubber.” The skeletal muscle of most marine mammals is very dark red, almost black, because of the relatively high myoglobin concentration. The design of the musculoskeletal system profoundly influences any mammal’s power output because it affects both thrust and propulsive efficiency (Pabst et al., 1999). Thrust forces depend on muscle morphology and the mechanical design of the skeletal system. The propulsive efficiency of the animal depends on the size, shape, position, and behavior of the appendage(s) used to produce thrust. Terrestrial mammals usually use their appendicular musculoskeletal system to swim using the proverbial dog paddle—alternate strokes of the forelimbs (and sometimes hind limbs). Pinnipeds use their more-derived appendicular musculoskeletal systems to swim. Unlike the other marine mammals, the fully aquatic sirenians and cetaceans swim using only their vertebral or axial musculoskeletal systems. Thus, in mammals that use their appendicular musculoskeletal systems to swim, two morphological “solutions” to increase thrust production are observed (Pabst et al., 1999). Proximal locomotor muscles tend to have large cross-sectional areas and so would have the potential to generate large in-forces. Proximal limb bones (i.e., humerus and femur) tend to be shorter than more distal bones (i.e., radius, ulna, tibia, and fibula), which increases the mechanical advantage of the lever system. The short proximal limb bones have an added hydromechanical benefit. These bones tend to be partially or completely enveloped in the body, which helps reduce drag on the appendage and increased body streamlining (Tarasoff, 1972; English, 1977; King, 1983). Contrast the distribution of muscle mass in the four species. Note that adaptations to each locomotory specialization have enlarged or reduced the corresponding muscles found in terrestrial mammals. Contrast the massiveness of the pectoral muscles (B-PEC) of the sea lion with those in the seal. The triceps (B-TRI) and deltoids (B-DEL) are also enlarged in both pinnipeds to increase thrust, and the olecranons (C,E-OLC) of both the seal and sea lion are enlarged to increase the mechanical advantage of these muscles. Note that the harbor seal has a unique component of the pectoral—an ascending pectoral muscle (B-PECa)—that extends over the humerus (also described for another phocid, the southern elephant seal; see Bryden, 1971). A dramatic change in thickness of the abdominal wall muscles (B-INT, EXT) occurs in young seals as they make the transition from a more terrestrial to a more aquatic lifestyle. Cetaceans and sirenians use their axial musculoskeletal systems to swim. Epaxial muscles (B-EPX) bend the vertebral column dorsally in upstroke; hypaxial muscles (B-HPX) and abdominal muscles bend the vertebral column ventrally in downstroke. Because there is no “recovery” phase, efficiency is increased. These muscles generate thrust forces that are delivered to the fluid medium via their flukes (Domning, 1977; 1978; Strickler, 1980; Pabst, 1990). The elongated neural spines (E-NSP) and transverse processes (E-TPR) of cetaceans also increase the mechanical advantage of the axial-muscle lever system, relative to that system in terrestrial mammals. By inserting far from the point of rotation, the lever arm-in is increased and, thus, force output is increased. A novel interaction between the tendons of the epaxial muscles and a connective tissue sheath that envelops those muscles also increases the work output of the axial musculoskeletal system in cetaceans (Pabst, 1993; Pabst et al., 1999). The *The term blubber is used differently in different species. In sea lions, seals, and manatees, it is subcutaneous fat in one or two layers, and resembles that found in terrestrial mammals. Blubber in cetaceans is fat—“inflated” dermis (Pabst et al., 1999).


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sirenian axial skeleton does not display elongated processes, which would increase the lever arm-in for dorsoventral flexion. Instead, the lumbar and cranialmost caudal vertebrae have elongated transverse processes (Domning, 1977; 1978).

The Diaphragm as a Separator of the Body Cavities The orientation of the diaphragm (C,D-DIA) in most marine mammals is very similar to the orientation of the diaphragm in the dog. Visualizing size, shape, and extent of the diaphragm will help one visualize the dynamics of respiration and diving. The diaphragm lies in a transverse plane and provides a musculotendinous sheet to separate the major organs of the digestive, excretory, and reproductive systems (all typically caudal to the diaphragm) from the heart with its major vessels; the lungs (C-LUN) and associated vessels and airways; the thyroid (C,D-THY), thymus (C,D-TYM), and a variety of lymph nodes, all located cranial to the diaphragm. The diaphragm is generally confluent with the transverse septum, so it attaches medially at its ventral extremity to the sternum. Although the diaphragm acts as a separator between the heart and lungs and the other organs of the body, the diaphragm is traversed by nerves and other structures, such as the aorta (D-AOR) (crossing in a dorsal and central position), the vena cava (D-CVC) (crossing more ventrally than the aorta, and often slightly left of the midline, although appearing to approximate the center of the liver), and the esophagus (D-EOS) (crossing slightly right of the midline, at roughly a midhorizontal level). This transverse orientation exists in most marine mammals, although the orientation of the diaphragm may be slightly diagonal, with the ventral portion more cranial than the dorsal portion. The West Indian manatee’s diaphragm differs from this general pattern of orientation and attachment. The manatee diaphragm and the transverse septum (D-TRS) are separate, with the latter occupying approximately the “typical” position of the diaphragm, and the diaphragm itself occupying a horizontal plane extending virtually the entire length of the body cavity. This apparently unique orientation presumably relates to buoyancy control (Rommel and Reynolds, 2000). There are two separate hemidiaphragms in the manatee. The central tendons firmly attach to hypapophyses (E-HYP) on the ventral aspects of the thoracic vertebrae, thereby producing the two pleural cavities.

Gross Anatomy of Structures Cranial to the Diaphragm Heart and Pericardium The pericardium is a fluid-filled sac surrounding the heart; in manatees, it often contains more fluid than is found in the typical mammal or in other marine mammals. The heart occupies a ventral position in the thorax (immediately dorsal to the sternum; D-STR). The heart lies immediately cranial to the central portion of the diaphragm (D-DIA; or the transverse septum in the manatee, D-TRS). In some species, the lungs (D-LUN) may embrace the caudal aspect of the heart, separating the caudal aspect of the heart from the diaphragm. As in all other mammals, marine mammal hearts have four chambers, separate routes for pulmonary and systemic circulation, and the usual arrangements of great vessels (venae cavae, D-CVC; aorta, D-AOR; coronary arteries; pulmonary arteries, PULaa; pulmonary veins, PULvv). Many marine mammal hearts are flattened from front to back (ventral to dorsal), are relatively squat from top to bottom, and have a rounded apex, giving them a shape quite different from the hearts of most terrestrial mammals (Drabek, 1975). Most pinnipeds and some cetaceans also have a distinctive dilatation of the aortic arch (Drabek, 1977). Cardiac fat occurs, but is rapidly lost in debilitated animals.



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Pleura and Lungs The pleural cavities and lungs (C-LUN) are generally found dorsal and lateral to the heart; in the manatee, the lungs are unusual in that they extend virtually the length of the body cavity and remain dorsal to the heart (Rommel and Reynolds, 2000). Lungs of some marine mammals (cetaceans and sirenians) are unlobed. The cranial ventral portion of the left lung in the bottlenose dolphin and other small odontocetes is very thin, almost veil-like, where it overlies the heart. Lobation in the pinnipeds is generally similar to that in the dog, that is, two lobes on the left (the cranial lobe has cranial and caudal parts) and three (including the accessory lobe) on the right. Reduction of lobation occurs in some phocids (Boyd, 1975; King, 1983). The terminal airways in all marine mammals are reinforced with either cartilage or muscle (Pabst et al., 1999). Apical (tracheal) bronchi are present in dolphins. In otariids, it is important to note that the bifurcation (D-BIF) of the trachea into the main-stem bronchi takes place at the thoracic inlet, not at the pulmonary hilus as is the case in phocids and cetaceans (McGrath et al., 1981; Nakakuki, 1993a,b; Wessels and Chase, 1998). The lungs of cetaceans are grossly smooth, but those of many pinnipeds are divided into distinct lobules in the ventral fields. Interestingly, sea otter lungs have distinct interlobular septa. The size of marine mammal lungs depends upon each species’ diving proficiency. Marine mammals that make deep and prolonged dives (e.g., elephant seals) tend to have smaller lungs than expected (based on allometric relationships), whereas shallow divers (e.g., sea otters) tend to have larger than expected lungs (Pabst et al., 1999).

Mediastinum The mediastinum is an artifact of the downward expansion of the lungs on either side of the heart in the typical mammal (Romer and Parsons, 1977); thus, the traditional definition of the mammalian mediastinum does not apply to manatees. The positions of the aortic hiatus, caval foramen (D-CAF), and esophageal hiatus (D-ESH) are unusual because of the configuration of the diaphragm. The manatee mediastinum (see manatee, layer D) is the midline region dorsal to where the pericardium attaches to the heart and ventral to the diaphragm, cranial to the transverse septum up to approximately the level of the first cervical vertebra. This is essentially what constitutes the cranial mediastinum of other mammals. The thyroid, thymus, tracheobronchial lymph nodes, and the tracheobronchial bifurcation are in the region defined as mediastinal in the manatee (Rommel and Reynolds, 2000). The mediastinum is thin and generally complete in the pinnipeds.

Thymus The thymus (C,D-TYM), which typically is relatively larger in young than in old individuals of any species, is found on the cranial aspect of the pericardium (sometimes extending caudally to embrace almost the entire heart) and may extend into the neck in otariids, the bottlenose dolphin (Cowan and Smith, 1999), and some other species.

Thyroids The thyroid glands (C,D-TYR) of the bottlenose dolphin and the manatee are located in the cranial part of the mediastinum along either side of the distal part of the trachea (C,D-TRA), prior to its bifurcation (D-BIF) into the bronchi. The paired, large, oval, dark-brown thyroid glands of pinnipeds, however, lie along the trachea just caudal to the larynx outside of the thoracic inlet (similar to the position in dogs).


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Parathyroids The parathyroid glands have been described in small cetaceans, and their location relative to the thyroid gland varies among species examined to date (Hayakawa et al., 1998). In Risso’s dolphins (Grampus griseus) they are dorsal to the thyroids or embedded within them, whereas in bottlenose dolphins they are on the surface of the thyroids and in the connective tissue surrounding the dorsal side of the thyroids. Little is known about the parathyroids of pinnipeds and sirenians.

Larynx The cetacean respiratory system has undergone several modifications that are associated with the production of sound. Immediately ventral and lateral to the blowhole (B,C,DBLO) are small sacs or lateral diverticulae (C-SAC). Medial to the diverticulae are the paired internal nares that extend on the cranial aspect of the braincase (D-BRN). The larynx (C-LAR), a spout-shaped structure referred to as the goosebeak, is composed of an elongated epiglottis and corniculate cartilage (Reidenberg and Laitman, 1987). The goosebeak extends through a small opening in the esophagus (supported laterally by an enlarged thyroid cartilage) into the relatively vertical narial passage; food can pass to either side of the goosebeak. Cetaceans have a robust hyoid apparatus (C,E-HYO) to support movements of the larynx. A palatopharyngeal sphincter muscle can keep the goosebeak firmly sealed (Pabst et al., 1999). For a detailed description of sound-producing anatomy, see Cranford et al. (1996).

Caval Sphincter One additional structure that is associated with the circulatory system, located on the cranial aspect of the diaphragm in seals and sea lions, is a feature atypical in mammals. This is the muscular caval sphincter (D-CAS), which can regulate the flow of oxygenated* blood in the large venous hepatic sinus (D-HPS) to the heart during dives (Elsner, 1969).

Microscopic Anatomy of Structures Cranial to the Diaphragm Respiratory System In cetaceans and otariids, cartilage extends around small bronchioles to the periphery of the lungs. In most phocids, cartilage is present around bronchi and bronchioles (Tarasoff and Kooyman, 1973; Boshier, 1974; Boyd, 1975). Bronchial glands are especially numerous in largercaliber bronchi and bronchioles of phocids. The configuration of terminal airways branching into alveoli varies among marine mammals, but, in general, respiratory ducts with small alveolar sacs make up the functional parenchyma. Myoelastic sphincters are present in the terminal bronchioles, presumably as an adaptation to diving (Boshier, 1974; Wessels and Chase, 1998). The number of alveolar duct units per lobule varies with species. The interalveolar septa have double rows of capillaries in most cetaceans and some otariids (e.g., in Steller but not California sea lions) but a single row of capillaries in phocids.

*In diving mammals with abundant arteriovenous anastomoses (shunts between arteries and veins before capillary beds), one can find high blood pressure and highly oxygenated blood in veins. One such venous reservoir of oxygenated venous blood is the hepatic sinus of seals (King, 1983).



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Thymus The thymus of marine mammals is composed of lobules, each with a distinct lymphocyte-rich cortex and a less cellular medulla. In many stranded immature marine mammals, there is profound thymic atrophy, with lymphoid depletion, and mineralization and keratinization of Hasell’s corpuscles.

Thyroids The thyroids of neonatal California sea lions, harbor seals, and elephant seals have plump cuboidal epithelium and little colloid (Little, 1991; Schumacher et al., 1993). In adults of the former two species, the epithelium also remains cuboidal, and the follicles remain fairly uniform in size. The thyroids of cetaceans are often distinctly lobulated, and the follicles of both young and adults are often small and lined with cuboidal epithelium similar to that of pinnipeds (Harrison, 1969b).

Parathyroids The parathyroids of Risso’s dolphins are divided into lobules by connective tissue, and have parenchymal cells consisting of chief cells with intracellular lipid droplets (Hayakawa et al., 1998).

Gross Anatomy of Structures Caudal to the Diaphragm Easy-to-find landmarks caudal to the diaphragm include a massive liver (C,D-LIV) and the various components of the gastrointestinal (GI) tract. The gonads and most other parts of the reproductive tracts are found only after the removal of the GI tract, except in a pregnant uterus.

Liver Typically, the liver is located immediately caudal to the diaphragm. It is a large, brownish, multilobed organ that tends to have most of its volume or mass positioned to the left of the body midline. Marine mammal livers are generally not too different from those of other mammals, although the manatee liver is a little more to the right and dorsal than are the livers of most other mammals. The number of lobes and the fissures in the lobes may vary, particularly in the sea lion’s liver, in which deep fissures give the lobes a deeply scalloped appearance. Bile may be stored in a gall bladder (often greenish in color) located ventrally, between lobes of the liver, although some mammals (e.g., cetaceans, horses, and rats) lack a gall bladder. Bile enters the duodenum (D-DUO) to facilitate chemical digestion of fats.

Digestive System Most of the volume of the cavity caudal to the diaphragm (the abdominal cavity) is occupied by the various components of the GI tract: the stomach, the small intestine (C-INTsml; duodenum, jejunum, ileum), and the large intestine (C-INTlg; cecum, colon, and rectum; C,D-REC). A strong sphincter marks the distal end of the stomach (the pylorus) before it connects with the small intestine (duodenal ampulla in cetaceans and sirenians). The separation between jejunum and ileum of the small intestine is difficult to distinguish grossly, although the two sections differ microscopically. The junction of the small and large intestines may be marked by the presence of a midgut cecum (homologous to the human appendix). The cecum is absent in most toothed whales, but present in some baleen whales (not the bowhead whale), vestigial but present in pinnipeds, and


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absent in sea otters. In manatees, the cecum is large, globular, and has two blind pouches called cecal horns. The large intestine, as its name implies, has a larger diameter than the small intestine in some marine mammals. In the sea lion, seal, and dolphin there is little difference in gross appearance between the small and large intestines. The cecum of sea lions and seals is about a meter from the anus, whereas the small intestines are about 20 times as long; in adult manatees, both the large and small intestines may approach or even exceed 20 m (Reynolds and Rommel, 1996). The proportions and functions of these components reflect feeding habits and trophic levels of the different marine mammals. Accessory organs of digestion include the salivary glands (C-SAL; absent in cetaceans, present in pinnipeds, very large in the manatee), pancreas (D-PAN), and liver. The pancreas is sometimes a little difficult to locate, because it can be a rather diffuse organ and decomposes rapidly; however, a clue to its location is its proximity to the initial part of the duodenum into which pancreatic enzymes flow (Erasmus and Van Aswegen, 1997). Another organ that is structurally, but not functionally, associated with the GI tract is the spleen (D-SPL), which is suspended by a ligament, generally from the greater curvature of the stomach in simple-stomached species, or from the first stomach in cetaceans). It is usually on the right side, but may have its greatest extent along the left side of the body. The spleen is usually a single organ, but in some species (mainly cetaceans), accessory spleens (occasionally referred to as hemal lymph nodes) may accompany it. It varies considerably in size among species; in manatees and cetaceans it is relatively small, but the spleen is relatively massive in some deep-diving pinnipeds (Zapol et al., 1979; Ponganis et al., 1992), where it acts to store red blood cells temporarily. The length and mass of the GI tract may be very impressive and create three-dimensional relationships that can be complex. Tough connective tissue sheets called mesenteries suspend the organs from the dorsal part of the abdominal cavity, and shorter connective tissue bands (ligaments*) hold organs close to one another in predictable arrangements (e.g., the spleen is almost always found along the greater curvature of the stomach and is connected to the stomach by the gastrosplenic ligament). Numerous lymph nodes and fat are also suspended in the mesenteries. The GI tracts of pinnipeds and other marine mammal carnivores follow the general patterns outlined above, although the intestines can be very long in some species (Schumacher et al., 1995; Stewardson et al., 1999). Cetaceans, however, have some unique specializations (Gaskin, 1978). In these animals, there are three or more compartments to the stomach, depending on the species. Functionally, the multiple compartments of cetacean stomachs correspond well to regions of the single stomach of most other mammals. Most cetaceans have three compartments; the first, called the forestomach (D-STM1; essentially an enlargement of the esophagus), is muscular and very distensible; it acts much like a bird crop (i.e., as a receiving chamber). The second (D-STM2), or glandular compartment, is the primary site of chemical breakdown among the stomach compartments; it contains the same types of enzymes and hydrochloric acid that characterize the “typical” mammalian stomach. Finally, the “U-shaped” third compartment, or pyloric stomach (D-STM3), ends in a strong sphincteric muscle that regulates flow of digesta into the duodenum of the small intestine. The initial part of the cetacean duodenum is expanded into a small saclike ampulla (occasionally mistaken for a fourth stomach). *Ligament has several meanings in anatomy: a musculoskeletal element (e.g., the anterior cuciate ligament), a vestige of a fetal artery or vein (e.g., the round ligament of the bladder), the margin of a fold in a mesentery (e.g., broad ligament), and a serosal fold between organs (e.g., the gastrolienal ligament). Note: In human terminology anterior and posterior are used; in comparative and veterinary terminology cranial and caudal are used when relating to the head and tail, respectively.



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Among the marine mammals, sirenians have the most remarkable development of the GI tract. Sirenians are herbivores and hindgut digesters (similar to horses and elephants), so the large intestine (specifically the colon) is extremely enlarged, enabling it to act as a fermentation vat (see Marsh et al., 1977; Reynolds and Rommel, 1996). The sirenian stomach is single chambered and has a prominent accessory secretory gland (the cardiac gland) extending prominently from the greater curvature. The duodenum is capacious and has two obvious diverticulae projecting from it. The GI tract of the manatee, with its contents, can account for more than 20% of an individual’s weight.

Urinary Tract The kidneys (C,D-KID) typically lie against the musculature of the back (B-HPX, hypaxial muscles), at or near the dorsal midline attachment of the diaphragm (crus, D-CRZ). In the manatee, the unusual placement of the diaphragm means that the kidneys lie against the diaphragm, not against hypaxial muscles. In many marine mammals, the kidneys are specialized as reniculate (multilobed) kidneys, where each lobe (renule) has all the components of a metanephric kidney. The reason that marine mammals possess reniculate kidneys is uncertain, but the fact that some large terrestrial mammals also possess reniculate kidneys has led to speculation that they are an adaptation associated simply with large body size (Vardy and Bryden, 1981), rather than for a marine lifestyle. Large body size may be important as the proximal convoluted tubules cannot be overlengthy and still conduct urine (Maluf and Gassmann, 1998). The kidneys are drained by separate ureters (D-URE), which carry urine to a medially and relatively ventrally positioned urinary bladder (C,D-BLD). The urinary bladder lies on the floor of the caudal abdominal cavity and, when distended, may extend as far forward as the umbilicus (A,B,C,D-UMB) in some species. The pelvic landmarks are less prominent in the fully aquatic mammals. In the manatee the bladder can be obscured by abdominal fat. Note that the renal arteries (D-REN) of cetaceans enter the cranial pole of the organ, and the ureters exit near the caudal pole, whereas in other marine mammals they enter and exit the hilus (typical of most mammals). Additionally, in manatees, there are accessory arteries on the surface of the kidney (Maluf, 1989).

Genital Tract Pabst et al. (1999) noted that the reproductive organs tend to reflect phylogeny more than adaptations to a particular niche. If one were to examine the ventral aspect prior to removal of the skin and other layers, one would discover that, especially in the sirenians and some cetaceans, positions of male and female genital openings are obviously different, permitting easy determination of sex without dissection. In all cases, the female urogenital opening (AU/G) is relatively caudal, compared with the opening for the penis in males. One way to approach dissection of the reproductive tracts is to follow structures into the abdomen from the external openings. The position and general form of the female reproductive tracts are similar to those of terrestrial mammals (Boyd et al., 1999). The vagina (C,D-VAG) opens cranial to the anus (A,B,C,D-ANS) and leads to the uterus (C,D-UTR), which is bicornuate in marine mammal species. The body of the uterus is found on the midline and is located dorsal to the urinary bladder (the ventral aspect of the uterus rests against the bladder). The uterine horns (cornua) extend from the uterine body toward the lateral aspects of the abdominal cavity. Implantation of the fertilized egg and subsequent placental development take place in the walls of the uterine horns, usually in the ipsilateral horn to ovulation (see Chapter 11, Reproduction). Dimensions of uterine horns vary with reproductive history and age. Often the fetus may expand the pregnant horn to occupy a substantial portion of the abdominal cavity. The horns terminate


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distally in an abrupt reduction in diameter and extend as uterine tubes (fallopian tubes) to paired ovaries (C,D-OVR). The uterus and ovaries are suspended from the dorsal abdominal wall by the broad ligaments. Uterine scars and ovarian structures may provide information about the reproductive history of the individual (Boyd et al., 1999; see Chapter 11, Reproduction). The ovaries of mature females may have one or more white or yellow-brown scars, called corpora albicantia and corpora lutea, respectively (see Chapter 11, Reproduction). Although ovaries are usually small solid organs, in sirenians they are relatively diffuse, with many follicles and more than one corpus albicans. The male reproductive tracts of marine mammals have the same fundamental components as those of “typical” mammals, but positional relationships may be significantly different. These differences are due to the testicond (ascrotal) position of the testes in many species (sea lion testes become scrotal when temperatures are elevated). The testes of some marine mammals are intra-abdominal* (DeSmet, 1977), whereas in phocids they are in the inguinal canal, covered by the oblique muscles and blubber (see Figure 2-20 in Pabst et al., 1999). The position of marine mammal testes creates certain thermal problems because spermatozoa do not survive well at body (core) temperatures; in some species, these problems are solved by circulatory adaptations mentioned below. The penis of marine mammals is retractable, and it normally lies within the body wall. General structure of the penis relates to phylogeny (Pabst et al., 1999). In cetaceans, it is fibroelastic type with a sigmoid flexure that is lost during erection, as seen in ruminants. Pinnipeds, sea otters and polar bears have a baculum within the penis, as do domestic dogs; in manatees it is muscular (see Chapter 11, Reproduction, and see Sexual Dimorphisms, below).

Adrenal Glands In marine mammals, adrenal glands (D-ADR) lie cranial to the kidneys and caudal to the diaphragm, as in terrestrial mammals. Adrenal glands can be confused with lymph nodes, but if one slices the organ in half, an adrenal gland is easy to distinguish grossly by its distinct cortex and medulla. In contrast, lymph nodes are more uniform in appearance.

Microscopic Anatomy of Structures Caudal to the Diaphragm Liver The histology of the liver of pinnipeds is quite similar to that of terrestrial mammals. In cetaceans, however, portal triads may have very thick-walled vessels (Hilton and Gaskin, 1978). Smooth muscle may also be found around some central veins (throttling veins) (Arey, 1941). Stainable iron (hemosiderosis) is common in neonatal harbor and northern elephant seals and in older otariids in captivity. Ito cells may be quite prominent in marine mammals, compatible with the presence of high vitamin A levels found in these livers (Rhodahl and Moore, 1943).

Digestive System The oropharynx of pinnipeds and odontocetes, and the caudal part of the odontocete tongue, are richly endowed with minor mucous glands, which enter out onto the mucosal surface via ducts that are visible grossly as small pits. Microscopically, the nonglandular and glandular stomachs resemble the analogous structures in terrestrial mammals. Parietal cells are exception*The position of the testes in sea otters is scrotal, and the testes of polar bears are seasonally scrotal (Reynolds et al., in press).



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ally prominent in odontocetes. In sirenians, the cardiac gland is a submucosal mass that protrudes cranially from the greater curvature of the stomach; it has a complicated folded lumen lined by mucous surface cells overlying long gastric glands lined with mucous and parietal cells. The glands of the main sac are lined by mucous cells and a lesser number of parietal cells (Marsh et al., 1977; Reynolds and Rommel, 1996). Histologically, the intestines of marine mammals are also similar to those of domestic mammals with the following exceptions (Schumacher et al., 1995). The villi are said to be absent in the proximal duodenum in some cetaceans, and Brunner’s glands are variably present. Plicae rather than villi are often present, creating chevron shapes on cross sections of cetacean intestine. The light and electron microscopic appearance of the small intestine of small odontocetes has been described in detail (Harrison et al., 1977). Gut-associated lymphoid aggregates are present throughout the intestines and may be diffuse or nodular. They are especially numerous in the distal colon of odontocetes and baleen whales, where they form the anal tonsil (Cowan and Brownell, 1974; Romano et al., 1993).

Urinary Tract Each reniculus has a histologically distinct cortex and medulla. Since cortex completely surrounds the medulla in the reniculi, ascending inflammation in one reniculus may spill over into the interstitium of an adjacent reniculus, giving the pattern of interstitial (hematogenous) nephritis. Thus, it is important to sample several reniculi from each kidney to assess pathological processes. In cetaceans there is normally a fibromuscular band at the corticomedullary junctions surrounding the medullary pyramid. Glomeruli of all species examined are of remarkably similar size (about one half the width of a 40× high dry field).

Genital Tract The morphology of the reproductive tract of the female varies with the stages of estrus and gestation (see Chapter 11, Reproduction). A description of cyclic changes in some of the cetaceans is given in Harrison (1969a) and in some sirenians in Boyd et al. (1999). Morphological changes of the genital mucosa associated with the estrous cycle have not been studied in detail in marine mammals, other than the harbor seal (Bigg and Fisher, 1974). In this species (described here to illustrate the variation in appearance through the estrous cycle), during follicular development then regression, the uterine mucosa increases in height and pseudostratification and then decreases to simple cuboidal. Uterine gland epithelium increases in height and secretory activity, and glands become increasingly coiled. Vaginal epithelium “destratifies” to become a “transitional-type” epithelium only a few cells thick, with vaginal pits (glands) lined by columnar epithelium with apical secretory product (goblet cell-like). The endometrial luminal and glandular epithelium of the nongravid horn is secretory and declines to cuboidal by parturition. During this luteal phase, there are subnuclear lipid vacuoles in the glandular epithelium. The vaginal epithelium is transitional during early placentation, but increases in secretory activity to become lined with tall columnar mucous cells with fingerlike projections of the lamina propria replacing the mucosal pits. During lactation, the morphology of both uterine and vaginal epithelium changes again. In the first part of lactation, the surface and glandular uterine epithelium is cuboidal, then undergoes hypertrophy and hyperplasia during the latter half of lactation. Luminal epithelium is occasionally pseudostratified, and the uterine stroma of both horns is edematous. The patchy hyperplasia and pseudostratification might be mistaken for dysplasia. Vaginal epithelium is almost transitional during the first part of lactation but proliferates to stratified squamous nonkeratinizing cells covered by sloughing mucous cells by the end of lactation. The endometrium of the gray seal prior to implanation is described by Boshier (1979; 1981).


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The placenta of pinnipeds is zonary, endotheliochorial, similar to that of domestic carnivores. In late gestation, it is often deep orange because of the marginal hematoma from which the fetus gains its iron stores in utero. After parturition and involution, old implantation sites may be visible grossly as dark areas in the mucosa, which are represented histologically by stromal hemosiderosis and arterial hyalinization. The placenta of cetaceans is diffuse epitheliochorial. The structure of the phocid corpus luteum is described by Sinha et al. (1972; 1977a). The prostate is the only accessory sex gland in pinnipeds and cetaceans (Harrison, 1969a). It is tubuloalveolar and has cuboidal to low-columnar to pseudostratified lining cells with basilar nuclei and pale apical cytoplasm. The fine structure of phocid testes and seminiferous tubules are described by Leatherland and Ronald (1979) and Sinha et al. (1977b), respectively.

Adrenals Pinniped adrenals may have an undulating or pseudolobulated cortex. In cetaceans, however, pseudolobulation is extensive and is created by connective tissue septae extending from the capsule. Large nerves, ganglia, and many blood vessels are associated with the hilus and capsular surface of pinniped adrenals.

Lymphoid and Hematopoietic Systems The capsules and trabeculae of pinniped lymph nodes are quite thick, and there is often abundant hilar and medullary connective tissue as well (Welsch, 1997). The degree of fibrosis seems to increase with age, and may be a function of chronic drainage reactions. Pinniped lymph nodes are organized like those of canids, having a peripheral subcapsular sinus, cortical follicular and interfollicular (paracortical) regions, and medullary cords and sinuses. Although some authors report that marine mammal lymphoid tissue is usually quiescent and lacks follicular development, secondary follicles are common in both peripheral and visceral lymph nodes of stranded pinnipeds, probably due to the common presence of skin wounds and visceral parasitism. In many stranded pinnipeds, the lymph nodes are sparsely but diffusely populated by lymphocytes, and the ghosts of germinal centers can be seen. Since this morphology is most common when the interval from death to post-mortem is prolonged, it has been interpreted to be a “washing out” of lymphocytes due to autolysis. The lymph nodes of some cetaceans are often deeply infolded or fused so that they appear to be organized similarly to the nodes of suids, whose follicular cortex is buried deep within the node and sinusoids and cords are located more toward the periphery. The correlation of anatomical location with nodal morphology has not been made for all species. The visceral nodes of the bottlenose dolphin have extensive smooth muscle in the capsule and trabeculae and have incomplete marginal sinuses (Cowan and Smith, 1999). The lymph nodes of the beluga are described by Romano et al. (1993). The elongated spleen of pinnipeds has a thick fibromuscular capsule and trabeculae with a sinusoidal pattern similar to that of canids. Periarteriolar reticular sheaths are more prominent in phocids than in otariids. The spherical spleen of cetaceans also has a thick capsule, which is fibrous externally and muscular internally, with the muscle cells extending into the thick trabeculae (Cowan and Smith, 1999). Extramedullar hematopoiesis is common in the spleens of pinniped and sea otter pups, but it seems to be uncommon in cetaceans.

Nervous System A detailed description of marine mammal neuroanatomy is beyond the scope of this chapter; for a comparison of some marine mammal brains (D-BRN), see Pabst et al. (1999). Suffice it



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to say that the brains of cetaceans and pinnipeds are large and well developed and have complex gyri in the cerebral and cerebellar cortices that are relatively larger than similarly sized brains of terrestrial mammals (Flanigan, 1972). The cetacean cerebrum is globoid and the rostral lobes extend ventrally. Like higher primates, cetaceans have well-developed temporal lobes (ventrolateral aspects of the cortices) that make brain removal a challenge. The pinniped brain is similar in orientation to the canine brain except for the larger cerebellum. In pinnipeds, the pineal gland is very large (up to 1.5 cm in diameter), especially in neonates (Bryden et al., 1986) and the size varies seasonally (see Chapter 10, Endocrinology). The pineal gland is located on the dorsal aspect of the diencephalon between the thalami and may be attached to the falx cerebri when the calvarium is removed at necropsy. There are no published descriptions of the pineal in cetaceans, and whether or not it exists is unclear. The pituitary gland is relatively large in both cetaceans and pinnipeds (Harrison, 1969b; Leatherland and Roland, 1976; 1978; Griffiths and Bryden, 1986). It is located within a shallow sella tunica in cetaceans and is surrounded by reams of blood vessels making it difficult to remove on necropsy. In pinnipeds, it is often sheared off during removal of the brain, so care should be taken to cut the lip of bone partially covering it to remove it intact. The spinal cord of phocids is relatively shorter than that of otariids; only the cauda equina occupies the lumbar and sacral canal. The cauda equina of the harbor seal pup is similar to that of the dog, but as they grow older, the cord changes significantly. The cauda equina starts in the lumbocaudal region in manatees. The region surrounding the cord—the vertebral canal—is significantly enlarged in seals, cetaceans, and sirenians. The neural canal is filled mostly with vascular tissue in seals and cetaceans and mostly with venous and fatty tissue in manatees. Manatee brains have pronounced lissencephaly and large lateral ventricles (Reep et al., 1989).

Circulatory Structures In general, blood vessels are named for the regions they feed or drain. Thus, the fully aquatic marine mammals (cetaceans and sirenians) lack femoral arteries, which supply the pelvic appendage. However, most organs in marine mammals are similar to those of terrestrial mammals, so their central blood supplies are also similar. The aorta (D-AOR) leaves the heart (D-HAR) as the ascending aorta, then forms the aortic arch (D-AAR) and roughly follows the vertebral column dorsal to the diaphragm as the thoracic aorta, which gives off segmental intercostal arteries and, in the case of cetaceans and manatees, feeds to the thoracic retia. Some of the segmental arteries of the dolphin anastomose at the base of the dorsal fin to form the single arteries that are arranged along the centerline of the dorsal fin (D-DFNaa). The aorta continues into the abdomen as the abdominal aorta, which gives off several paired (e.g., renal, gonadal) and unpaired (e.g., celiac, mesenteric) arteries. The caudal aorta follows the ventral aspect of the vertebrae in the tail; in the permanently aquatic marine mammals the caudal vessels are large when compared with the vessels in species with small tails. In the dolphin, the caudal arteries branch into dorsal and ventral superficial arrays of arteries (D-FLKaa; Elsner et al., 1974). In the permanently aquatic marine mammals, there are robust ventral chevron bones that form a canal in which the caudal aorta, its branches, and some veins (the caudal vascular bundle, D-CVB) are protected. This site is convenient in some species for venipuncture; however, note that it is an arteriovenous plexus, so samples collected may be mixed arterial and venous blood. Some of the diving mammals (e.g., seals, cetaceans, and sirenians) have few or no valves in their veins (Rommel et al., 1995); this adaptation simplifies blood collection because the blood can drain toward the site from both directions, although blood collection is complicated by the arteriovenous plexuses described above. Other exceptions to the general pattern of mammalian


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circulation are associated with thermoregulation and diving. Countercurrent heat exchangers abound, and extensive arteriovenous anastomoses exist to permit two general objectives to be fulfilled: (1) regulating loss of heat to the external environment while keeping core temperatures high, and (2) permitting cool blood to reach specific organs (e.g., testes and epididymides, ovaries and uteri) that cannot sustain exposure to high body temperatures (see reviews by Rommel et al., 1998; Pabst et al., 1999). Mammals have three options for blood supply to the brain: the internal carotid, the external carotid, and the vertebral arteries. Some species use only one and others two, but the manatees use all three pathways. Cetaceans have a unique blood supply to the brain (D-BRN); the blood to the brain first enters the thoracic retia, a plexus of convoluted arteries in the dorsal thorax. Blood leaves the thoracic retia and enters the spinal retia, where it surrounds the spinal cord and enters the foramen magnum (McFarland et al., 1979). There are two working hypotheses for this convoluted path to the brain: (1) the elasticity of the retial system allows mechanical damping of the blood pulse pressure wave (McFarland et al., 1979; Shadwick and Gosline, 1994), and (2) the juxtaposition of the thoracic retia to the dorsal aspect of the lungs may provide thermal control of blood entering the spinal retia (Rommel et al., 1993b). Combined with cooled blood in the epidural veins, the spinal retia may provide some temperature control of the central nervous system (Rommel et al., 1993b). Carotid bodies, important in regulation of blood flow, have been documented in the harbor seal (Clarke et al., 1986).

The Potential for Thermal Insult to Reproductive Organs Mammals maintain high and, in most species, relatively uniform core temperatures. Because they live in water, which conducts heat 25 times faster than air at the same temperature, many marine mammals have elevated metabolic rates and/or adaptations to reduce heat loss to the environment (Kooyman et al., 1981; Costa and Williams, 1999). Aquatic mammals with low metabolic rates must live in warm water or possess even more elaborate heat-conserving structures. Most mammalian tissues tolerate limited fluctuations in temperature, and some tissues, such as muscle, perform better at somewhat higher temperatures. However, reproductive tissues are particularly susceptible to thermal insult, and various mechanisms have evolved to protect them (VanDemark and Free, 1970; Blumberg and Moltz, 1988). In terrestrial mammals, production and storage of viable sperm requires a relatively narrow range of temperatures. Temperatures between 35 and 38°C can effectively block spermatogenesis (Cowles, 1958; 1965). Abdominal temperatures can detrimentally affect long-term storage of spermatozoa in the epididymides in many species (Bedford, 1977). In many mammals, the scrotum provides a cooler environment by allowing the sperm-producing tissues to be positioned outside the abdominal cavity, away from relatively high core temperatures. Additionally, in scrotal mammals, the pampiniform plexus can, via countercurrent heat exchange, reduce the temperature of arterial blood from the core to the testes and help keep testicular temperature below that of the core (Evans, 1993). The skin of the scrotum is well vascularized, has an abundance of sweat glands, and is highly innervated with temperature receptors. Muscles in the scrotal wall involuntarily contract and relax in response to cold and hot temperatures, respectively. The exposed scrotum provides a thermal window through which heat may be transferred to the environment, thereby regulating the temperature of sperm-producing tissues. Interestingly, the morphological adaptations for streamlining observed in some marine mammals create potentially threatening thermal conditions for the reproductive systems of diving mammals. The primary locomotory muscles of terrestrial mammals are appendicular,



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so much of the locomotory heat energy of the muscle is transferred to the environment rather than directed into the body cavities; this is not the case for ascrotal marine mammals, whose primary locomotory muscles surround the abdominal and pelvic cavities. A factor that may increase core temperature of marine mammals is change in blood flow patterns during diving. Marine mammals can dramatically redistribute their cardiac output during dives, resulting in severely reduced blood flow to some body tissues, such as muscles and viscera (Elsner and Gooden, 1983; Kooyman, 1985). In terrestrial mammals, redistributions of cardiac output in response to physiological conditions such as exercise, feeding, thermoregulation, and pregnancy are relatively well known (Elsner, 1969; Baker and Chapman, 1977; Baker, 1982; Blumberg and Moltz, 1988). For example, in humans, large increases in muscle temperature (as high as 1°C/min) have been measured during the ischemia at the onset of exercise (Saltin et al., 1968). Surprisingly, the magnitude of routine cardiovascular adjustments undergone by marine mammals during prolonged dives (Elsner, 1999) is approached in terrestrial mammals only during pathological conditions such as hyperthermia and hypovolemic shock. The axial locomotion of pinnipeds, cetaceans, and manatees requires a relatively large thermogenic muscle mass around the vertebral column and abdominal organs. Blubber insulates these thermogenic muscles, suggesting the potential for elevated temperatures at the reproductive systems, particularly during the ischemia of prolonged dives. The temporary absence of cooling blood through locomotory muscles increases the probability of severe thermal consequences for the diving mammal. Abdominal, or partly descended, testes (cryptorchidism) result in sterility in many domestic mammals and humans. Ascrotal testes are typical for many marine mammals, such as phocid seals, dolphins, and manatees. There are vascular adaptations that prevent deep-body hyperthermic insult in cetaceans and phocids (Rommel et al., 1998). In dolphins, cooled venous blood is delivered to an inguinal countercurrent heat exchanger to cool the testes and epididymides indirectly, whereas, in phocid seals, cooled venous blood is delivered to an inguinal venous plexus to cool the testes and epididymides directly. Similar structures prevent reproductive hyperthermic insult in females (Rommel et al., 1995). One additional vascular adaptation that may have significant influence on diving is the presence of cooled blood in the large vascular structures within the vertebral canal, adjacent to the spinal cord. The large epidural veins (dolphins, seals, and manatees) and spinal retia (dolphins) may influence spinal cord temperature and, thus, influence dive capabilities, by modifying regional metabolic rates (Rommel et al., 1993b). The central nervous system is temperature sensitive, and lowering cord temperature influences global metabolic responses.

Skeleton Knowledge of the skeleton offers landmarks for soft tissue collection and provides an estimate of body size from partial remains (Rommel and Reynolds, in press). Traditionally, the postcranial skeleton is subdivided into axial components (the vertebral column, ribs, and sternabrae, which are “on” the midline) and appendicular components (the forelimbs, hind limbs, and pelvic girdle, which are “off ” the midline). The scapulae and humeri of the forelimbs are indirectly attached to the body, essentially by tensile elements (muscles and tendons); in contrast, the hind limbs are attached via a pelvis directly to the vertebral column and thus are able to transmit both tension and compression to the body. The skeleton supports and protects soft tissues, controls modes of locomotion, and determines overall body size and shape; the marrow of some bones may generate the precursors of certain blood cells. While the animal is alive, bones are continuously remodeled in response to biochemical and biomechanical demands and, thus, offer information that can help


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biologists interpret events in the life history of the animal after its death. Skeletal elements contribute to fat (particularly in the cetaceans) and calcium (particularly in the sirenians) storage and thus influence buoyancy. The sea lion propels itself through the water by its forelimbs, and its skeletal components are relatively massive in that region. On land, its forelimbs can act as fulcra for shifting the center of mass by changing the shape of its neck and the trunk (for more, see English, 1976a,b; 1977). The permanently aquatic species locomote with a dorsoventral motion of the trunk and elongated tail. This dorsoventral motion of the axial skeleton is characteristic of almost all mammalian locomotion. In contrast, the seal uses lateral undulations of its trunk and hind flippers when swimming (like a fish), yet it may locomote on land with dorsoventral undulations, like its terrestrial ancestors. Relative motion between vertebrae is controlled, in part, by the size and shape of the intervertebral disks. The intervertebral disks resist the compression that skeletal muscles exert and tend to force vertebrae together. Intervertebral disks are composite structures, with a fibrous outer ring, the annulus fibrosus, and a semiliquid inner mass, the nucleus pulposus. The outermost fibers of the annulus are continuous with the fibers of the periosteum. The flexibility of the vertebral column depends, in part, on the thickness of the disks. Intervertebral disks are a substantial proportion (10 to 30%) of the length of the postcranial vertebral column. The intervertebral disks provide flexibility but are not “responsible” for the general curvature of the spine—the nonparallel vertebral body faces provide the spinal curvature. For convenience, the vertebral column is separated into five regions, each of which is defined by what is or is not attached to the vertebrae. These regions are cervical, thoracic, lumbar, sacral, and caudal. In some species, the distinctions between vertebrae from each region are unambiguous. However, in some other species the distinctions between adjacent regions are less obvious. This is particularly true in the permanently aquatic species, where there is little or no direct connection between the pelvic vestiges and the vertebral column. The vertebral formula varies within, as well as among, species. The number of vertebrae, excluding the caudal vertebrae, is surprisingly close to 30 in most mammals (Flower, 1885). Most mammals have seven cervical, or neck, vertebrae (sirenians and two-toed sloth have six and the three-toed sloth has nine), whereas the number of thoracic and lumbar vertebrae varies between species. The number of sacral vertebrae is commonly two to five, but there are exceptions. The number of caudal vertebrae varies widely—long tails usually have numerous caudal vertebrae. The cervical vertebrae are located cranial to the rib-bearing vertebrae of the thorax. Some cervical vertebrae have movable lateral processes known as cervical ribs, none of which makes contact with the sternum. Typically, the permanently aquatic marine mammals have short necks, even if they have seven cervical vertebrae. However, the external appearance of a short neck in seals is misleading. Close comparison of the seal and sea lion skeletons reveals that they have quite similar neck lengths, although the distribution of body mass is different. Seals often hold their heads close to the thorax, which causes a deep “S” curve in the neck. This provides the seals with a “slingshot potential” for grasping prey (or careless handlers). The shapes of the seal neck vertebrae are complex to allow this curve. Serial fusion (ankylosis) of two or more cervical vertebrae is common in the cetaceans, although in some cetaceans (e.g., the narwhal, beluga, and river dolphins), all the cervical vertebrae are unfused and provide considerable neck mobility. The rib-bearing vertebrae are the thoracic vertebrae, and the thoracic region is defined by the presence of movable ribs. The authors distinguish between vertebral ribs (E-VBR), which are associated with the vertebrae, and “sternal ribs” (E-SBR), which are associated with the sternum. This distinction is made because some odontocetes, unlike most other mammals,



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have bony rather than cartilaginous sternal ribs (bony “sternal ribs” are also found in the armadillo). “Costal cartilages” is an acceptable alternative term for sternal ribs if the sternal ribs are never ossified (calcification with old age does not count). Some thoracic vertebrae have ventral vertebral projections called hypapophyses (see the manatee, E-HYP)—not to be confused with chevron bones, which are intervertebral and not part of the caudal vertebrae. In the manatee, the diaphragm is firmly attached along the midline of the central tendon to hypapophyses. Hypapophyses also occur in some cetaceans (e.g., the pygmy and dwarf sperm whales, Kogia) in the caudal thorax and cranial lumbar regions. It is assumed that these hypapophyses increase the mechanical advantage of the hypaxial muscles much as do the chevrons (Rommel, 1990). The neural spines (E-NSP) of thoracic vertebrae of many mammals are often longer than those in any other region of the body. Long neural spines provide mechanical advantage to neck muscles that support a head cantilevered in front of the body. Terrestrial species with large heads tend to have long neural spines, but in aquatic mammals the buoyancy of water negates this reason for long neural spines.

Ribs Embryologically, ribs and transverse processes develop from the same precursors. Thus, some aspects of ribs are similar to those of transverse processes (E-TPR). It is the formation of a movable joint that distinguishes a rib from a transverse process. An unfinished joint may be indicative of developmental age. In some species (i.e., the manatee) there may be a movable “rib” (pleurapophysis) on one side and an attached “transverse process” on the other side of the same (typically the last thoracic) vertebra (Rommel and Reynolds, 2000). Ribs may attach to their respective vertebrae at one or more locations (e.g., centrum, transverse process). Typically, the cranialmost ribs have two distinct regions of articulation (capitulum and tuberculum) with juxtaposed vertebrae and are referred to as double headed. The caudalmost ribs have single attachments and are referred to as single headed. In most mammals, the single-headed ribs have lost their tubercula and are attached to their vertebrae at the capitulum on the centrum. In contrast, the single-headed ribs of cetaceans lose their capitula and are attached to their respective vertebrae by their tubercula on the transverse processes (Rommel, 1990). The last ribs (E-LRB) often “float” free from attachment at one or both ends; these ribs tend to be significantly smaller than the ones cranial to them, and they are often lost in preparation of the skeleton. The ribs of some marine mammals are more flexible than those of their terrestrial counterparts; this flexibility is an adaptation to facilitate diving. Ribs are illustrated in layer E in the correct posture for a healthy animal. Note that all illustrated species but the manatees have oblique angles between the rib shaft and the long axis of the body. As the hydraulic pressures increase with depth, the ribs rotate to avoid bending with changes in thoracic cavity volume.

Sternum The sternum (D,E-STR) is formed from bilaterally paired, serial elements called sternabrae. The paired elements fuse on the midline, occasionally imperfectly, leaving foramina in the sternum. The cranialmost sternal ribs (E-SRB, also called costal cartilages) extend from the vertebral ribs to articulate firmly with the sternum at the junctions between sternebrae. The first sternal rib articulates with the manubrium (C,D-MAN) cranial to the first intersternabral joint. The manubrium may have an elongate cartilaginous extension (e.g., in seals), and the first sternal rib is often different from the more caudal sternal ribs (typically larger and more robust). In some mysticetes, only the manubrium is formed, and only the first rib has a bony attachment to it. The subsequent ribs articulate with a massive cartilaginous structure that extends from the caudal


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aspect of the manubrium (which may be referred to as a pseudosternum). The xiphoid process (E-XIP, last sternabra) is also different; it too may articulate with more than one (often many) sternal rib(s) and have a large cartilaginous extension.

Postthoracic Vertebrae Some authors avoid the difficulties of defining the lumbar, sacral, and caudal regions in the permanently aquatic species by lumping them into one category—the postthoracic vertebrae; by “lumping,” these authors avoid some interesting comparisons. Generally, the lumbar vertebrae are trunk vertebrae that do not bear ribs, and the number of lumbar vertebrae is closely linked to the number of thoracic vertebrae, but not always. Note that the caudal vertebrae of cetaceans start with the start of the chevron bones, and extend to the tip of the tail (fluke notch, A-NOC), whereas manatee vertebrae stop 3 to 9% of the total body length (as much as 17 cm in a large specimen) from the fluke tip (E-LVR).

Sacral Vertebrae There are at least two commonly accepted definitions for sacral vertebrae: (1) serial fusion of postlumbar vertebrae, only some of which may attach to the pelvis (the human os sacrum), and (2) only those that attach to the ilium, whether or not they are serially fused. Both definitions have merit. Within species, the number of serially ankylosed vertebrae may vary, particularly with age. Additional landmarks are the exit of spinal nerves from the neural canal and the foramina for segmental blood vessels. In species with a bony attachment between the vertebral column and the pelvis, the definition of sacral is easy. However, in the cetaceans and some sirenians (dugongs have a ligamentous attachment between the vertebral column and the pelvic vestiges), there are no sacral vertebrae by definition.

Chevron Bones The chevron bones are ventral intervertebral ossifications in the caudal region. By definition, each is associated with the vertebra cranial to it (note that there is some controversy over which is the first caudal vertebra; see Rommel, 1990). Chevron bone pairs are juxtaposed (in manatees) or fused (in dolphins, but not always) at their ventral apexes and articulate dorsally with the vertebral column to form a triangular channel. Within the channel (hemal canal) are found the blood vessels to and from the tail. In some species, the ventral aspects of each chevron bone fuse and may continue as a robust ventral protection that can function to increase the mechanical advantage of the hypaxial muscles to ventroflex the tail. In some individuals, the first two or three chevrons may remain open ventrally but fuse serially on either side.

Pectoral Limb Complex The forelimb skeleton includes the scapula, humerus, radius and ulna, and manus. The scapula is attached to the axial skeleton only by muscles. There is no functional clavicle in marine mammals (Strickler, 1978; Klima et al., 1980). The scapula consists of an essentially flat (slightly concave medially) blade with an elongate scapular spine extending laterally from it. The distal tip of the spine, if present, is the acromion. The scapular spine is roughly in the center of the scapular blade in most mammals. However, in cetaceans, the scapular spine is close to the cranial margin of the scapular blade, and both the acromion and coracoid extend beyond the leading edge of the blade. The humerus (E-HUM) has a ball-and-socket articulation in the glenoid fossa of the scapula— this is a very flexible joint. The humerus articulates distally with the radius (E-RAD) and ulna (E-ULN); this is also a flexible joint in most other mammals, but it is constrained in cetaceans. The olecranon is a proximal extension of the ulna that increases the mechanical advantage of the



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triceps muscles that extend the forelimb. In species like the sea lion, the olecranon is robust; however, in the cetacea, it is relatively small. The radius and ulna of manatees fuse at both ends as the animal ages. This fusion prevents axial twists that pronate and supinate the manus. The radius and ulna of cetaceans are also constrained but not typically fused. The distal radius and ulna articulate with the proximal aspect of the manus. The manus includes the carpals, metacarpals, and phalanges (English, 1976). There are five “columns” of phalanges, each of which is called a digit. The digits are numbered starting from the cranial aspect (the thumb, which is digit one, associated with the radius). In many of the marine mammals, the “long” bones of the pectoral limb (humerus, radius, and ulna) are relatively short, and the phalanges are elongated. Cetaceans are unique among mammals in that they have more than the maximum number of phalanges found in all other mammals; this condition is known as hyperphalangy (Howell, 1930). The number varies within each species—the bottlenose dolphin has a maximum number of nine digits.

Pelvic Limb Complex The typical mammalian pelvis is made of bilaterally paired bones: ilium, ischium, pubis, and acetabular bone (the paired ossa coxarum), one to three caudal vertebrae, and the sacrum. Each of the halves of the pelvis attaches (via the ilium) to one or more sacral vertebrae. The crest of the ilium (C,E-ILC) is a prominent landmark that flares forward and outward beyond the region of attachment between the sacrum and the ilium. The ossa coxarum join ventrally along the midline at the pelvic symphysis, which incorporates the pubic bone cranially and the ischiatic bone caudally. In the permanently aquatic marine mammals, there is but a vestige of a pelvis (E-PEL) to which portions of the rectus abdominis muscles (B-REC) may attach. Additionally, the crura of the penis may be supported by these vestiges (Fagone et al., 2000). In some of the large whales, there is occasionally a vestige of a hind limb articulating with the pelvic vestige. The hind limb, if present, articulates with the vertebral column via a ball-and-socket joint at the hip. The proximal limb bone is the femur (C,E-FEM). The socket of the pelvis, the acetabulum, receives the head of the femur. Distally, the femur articulates with the tibia and the fibula (as the stifle joint). The tibia and fibula distally articulate with the pes, or foot. The pes is composed of the tarsals proximally, the metatarsals, and the phalanges distally. Note that the digits of the sea lion terminate a significant distance from the tips of the flipper.

Sexual Dimorphisms In many mammals, the adult males are larger than the adult females. In marine mammals, this size difference is at its extreme in otariids, elephant seals, and the sperm whales. In contrast, the adult females of the baleen whales and some other species are larger than the adult males. In the permanently aquatic marine mammals, there may be sexual dimorphisms in the pelvic vestiges (Fagone et al., 2000). The penises of mammals are supported by crura consisting of a tough outer component (tunica albuginea) and the cavernous erectile central component (corpus cavernosum), which attach to the ischiatic bones of the pelvis. The muscles that engorge the penis with blood are also attached to the pelvis. Presumably, the mechanical forces associated with these muscles influence pelvic vestige size and shape, particularly in manatees. Males in some groups of mammals, particularly the carnivores, have a bone within the penis (the baculum) that helps support the penis. Growth rate of the os penis differs from that of the appendicular skeleton in some species (Miller et al., 1998).


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Bone Marrow Bone marrow of cetceans is vertebral as well as costal. Because the marrow cavity of the bones of marine mammals generally retains abundant trabecular bone throughout life, it is best to examine the marrow histologically via impression smears of cut surface or in decalcified sections. Most manatee bones are amedullary (Fawcett, 1942), so usable marrow impression smears are restricted to vertebrae.

Acknowledgments The authors thank Meghan Bolen, Judy Leiby, James Quinn, John Reynolds, Lisa Johnson, and Terry Spraker for reviewing the manuscript, Dan Cowan for information on parathyroids, and Frances Gulland and Rebecca Duerr at The Marine Mammal Center for helpful discussions. Anatomical illustrations were created with FastCAD (Evolution Computing, Tempe, AZ).

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10 Endocrinology David J. St. Aubin

Introduction The biochemicals classified as hormones are exceedingly potent agents, capable of profoundly influencing cellular functions to establish the optimum internal environment for a particular set of environmental challenges or survival needs. By definition, these chemicals are produced within well-defined glands or organs, secreted into blood or other extracellular media, and transported at least some distance to exert their effects on unrelated tissues. Endocrine systems are typically regulated through stimulatory and negative feedback mechanisms, often involving separate endocrine glands in a cascading sequence of hormone release originating from central neurological structures. Other biochemical stimuli, such as rising blood glucose or changes in the ratio of sodium to potassium (Na:K) in plasma, are equally capable of eliciting endocrine responses from the structures that are responsible for maintaining those constituents within appropriate physiological limits. The basic principles of vertebrate endocrinology, as presented in recent reference publications (Wilson et al., 1998), appear to hold for marine mammals. There are, nevertheless, some interesting adaptations, driven by the peculiar life histories of these animals, that represent important deviations from the norm for terrestrial mammals and need to be taken into account by both the researcher and the clinician. Some of these endocrine systems have received considerable attention in the literature, as extensively reviewed by Kirby (1990); for others, the available information is scant and deserves the attention of marine mammal physiologists and endocrinologists. The considerable and growing body of data on reproductive endocrinology will be examined in a separate chapter (see Chapter 11, Reproduction) focused on that specific aspect of marine mammal biology. Information on the status and role of various endocrine systems is invaluable to those seeking to understand better how marine mammals are able to survive the rigors of a most challenging environment. Prolonged fasts, deep dives, seasonally synchronized molting and breeding cycles, and an osmotically hostile medium, all require a metabolism finely tuned by endocrine controls. Breakdowns in these systems can significantly compromise the health and survival of the organism. The activity of specific endocrine organs, as measured by hormone levels in body fluids and excretions, can provide important information about the internal environment of the subject, and guide corrective therapy. Although large, the body of information on marine mammal endocrinology holds little regarding primary endocrinopathies, when compared with terrestrial mammals. More often, endocrine imbalances in marine mammals reflect perturbations in other systems, and the challenge is not only to establish what the primary cause might be, but also to recognize what physiological changes might be attributable to the secondary endocrine dysfunction. 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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Sample Collection and Handling Blood The most commonly collected specimen for hormonal analysis is blood. Serum is preferred for most analyses, particularly for those in which anticoagulants have been identified as interfering with results. According to one manufacturer of radioimmunoassays (RIA) (Diagnostic Products Corporation, Los Angeles, CA), heparinized plasma yields satisfactory results, except for the measurement of free triiodothyronine (f T3), whereas EDTA-treated blood is generally unsuitable. Fasting is not usually a prerequisite for obtaining a sample for thyroid and adrenal hormone analysis, but highly lipemic samples collected during the absorptive phase after eating are unsuitable for thyroid hormone (TH) testing. For hormones such as cortisol, known to exhibit diurnal variation, it is important to standardize, or at least note, the time of day at which the specimen is collected to interpret the results properly. Most hormones, particularly steroids and TH, are quite stable in serum samples refrigerated for 2 to 3 days or stored frozen at −70°C for months. Thawed samples should not be refrozen.

Saliva The measurement of hormones in saliva represents an attractive alternative as a noninvasive technique (Theodorou and Atkinson, 1998). Nevertheless, its collection requires either wellestablished behavioral control or full restraint, either of which can be used for the collection of blood samples. Laboratories are becoming better equipped to test saliva, and this is likely to result in more extensive reference data and established correlations with circulating levels of the hormone in question. One manufacturer of testing kits (Salimetrics LLC, State College, PA) recommends the use of plain, non-citric acid–treated, cotton Salivettes® (Sarstedt, Leicester, UK). Saliva samples should be frozen prior to assay to precipitate mucins. The approach has been investigated for monitoring reproductive hormones in marine mammals (Theodorou and Atkinson, 1998) (see Chapter 11, Reproduction), but there are insufficient data on other endocrine systems to establish its utility at this time.

Feces Fecal analysis of corticosteroids and reproductive hormones has proved useful in monitoring the endocrine status of terrestrial mammals (Brown et al., 1994), and has been attempted in at least one study on cortisol in harbor seals (Phoca vitulina) (Gulland et al., 1999). Samples may be frozen for months prior to analysis. Cortisol was extracted in a solution of buffered saline and 50% ethanol containing 0.1% bovine serum albumin and 5% Tween 20 (Zymed Laboratories, Inc., San Francisco, CA), and then assayed using conventional radioimmunoassay techniques. Cortisol concentrations up to 1100 µg/kg were reported, but could not be correlated with plasma values obtained either at the approximate time of fecal collection or at the peak of adrenocortical stimulation on the previous day. Further studies are needed to allow the use and interpretation of fecal hormone data for marine mammals.

Urine Hormones responsible for fluid and electrolyte balance, such as aldosterone and vasopressin, have been analyzed in urine samples of phocid seals (Hong et al., 1982). A 24-hour sample is optimal to integrate the daily fluctuations associated with consumption of food and water, which presents some impediment to investigations in marine mammals that cannot be confined or held out of water for the duration. Behavioral collection of urine has been established in


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cetaceans, but still cannot ensure that some of the daily urine production has not been lost into the environment. Samples for aldosterone determination should be refrigerated during or immediately after collection, and are stabilized with 1 g of boric acid/100 ml; they may be refrigerated for up to a week or stored frozen at −20°C for a month. No preservative is required for cortisol.

Tissues Palmer and Atkinson (1998) established a methodology for analyzing the corticosteroid content of blubber biopsies, specimens that are routinely collected for genetic studies on free-ranging cetaceans, particularly large whales from which blood, saliva, and urine are virtually impossible to acquire. Once validated, relative to more established measures of circulating hormone concentrations, the approach could prove useful in field studies on mysticetes, among others.

Pineal Gland Marine mammals exhibit strong seasonality in activities such as reproduction and molt. Synchronization of such events with appropriate environmental conditions is critical to optimizing survival, and likely requires the ability to sense cues that signal important seasonal events. Changes in air and water temperatures and daylength, particularly at midtemperate to high latitudes, can be pronounced enough to trigger significant annual events, such as migration in humpback whales (Megaptera novaeangliae) (Dawbin, 1966) (see Chapter 1, Sentinels). The hormone melatonin is considered to play a critical role in the integration of endocrine physiological systems with photoperiod in mammals (Goldman, 1983; Vivien-Roels and Pévet, 1983). Although at present of minimal clinical significance in marine mammals, the sporadic research that has been undertaken, particularly in pinnipeds, has identified the critical role of melatonin in early metabolism and subsequent seasonal activities. The principal source of melatonin is the pineal gland (epiphysis) typically located above the third ventricle of the brain. Other tissues, such as the retina, intestines, red blood cells, and salivary glands, contribute to circulating levels, and may represent significant sources in cetaceans, for which the very existence of a discrete pineal has been controversial (Flanigan, 1972). Nevertheless, Arvy (1970) and Behrmann (1990) have described the organ in several species of small odontocetes. This contrasts to the prominence of the gland in some pinnipeds, notably the Weddell seal (Leptonychotes weddellii) (Cuelo and Tramezzani, 1969; Bryden et al., 1986), northern fur seal (Callorhinus ursinus) (Elden et al., 1971), and northern (Mirounga angustirostris) (Bryden et al., 1994) and southern elephant seals (M. leonina) (Bryden et al., 1986; Little and Bryden, 1990). Earlier work on northern fur seals had recognized the pineal’s impressive dimensions and activity relative to those in humans, and suggested that further investigation might provide useful insights into the physiological role of melatonin in mammals (Elden et al., 1971). Weighing as much as 9 g in the newborn southern elephant seal (Little and Bryden, 1990), the gland can be roughly the size of the entire brain of a hamster, the species that has contributed most substantially to the understanding of melatonin physiology (Goldman, 1983). Elephant seals continue to show substantial changes in the size of their pineal throughout life. The gland is largest in the dark of winter, weighing up to 2 g/1000 kg of body weight, and regresses to less than half of that in nearly constant daylight in the summer (Griffiths et al., 1979; Griffiths and Bryden, 1981; Griffiths, 1985). No less remarkable are the fluctuations in circulating concentrations of melatonin that are most evident soon after birth in southern elephant seals (Table 1). Levels approaching 69,000 pg/ml have been recorded in neonates (Little and Bryden, 1990), with concentrations diminishing to


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TABLE 1 Reported Concentrations (pg/ml) of Melatonin in Pinnipeds Species Cystophora cristata (hooded seal) Halichoerus grypus (gray seal) Leptonychotes weddellii (Weddell seal)

Mirounga angustirostris (northern elephant seal)

Mirounga leonina (southern elephant seal)

Pagophilus groenlandicus (harp seal)

Specimens Neonate, 24-h sample Neonate, 24-h sample Pup (4 d), 24-h sample Pup (10 d), 24-h sample Pup (0–10 d) day Pup (12–35 d) day Juvenile (60 d) Adult Pup (0–5 d) day Pup (0–5 d) night Pup (6–25 d) day Pup (4 wk) night Pup (4 wk) day Juvenile (10 wk) Adult Neonate 0–24 h Pup (0–5 d) Pup (6–20 d) Juvenile Postpubertal Adult Neonate (1–2 d) Pup (2 wk)

Melatonin (pg/ml) 0–6000 100–7000 0–3000 0–450 50–>1000 50–220 53 5–12 695–1159 1200–>2318 2 wk) Adults

2.1–2.3 1.3–2.7

Adults and juveniles, molting Juveniles, both sexes

4.0

Stokkan et al., 1995; Woldstad et al., 1999 Engelhardt and Ferguson, 1980; Stokkan et al., 1995; Hall et al., 1998; Woldstad et al., 1999 Boily, 1996; Hall et al., 1998 Engelhardt and Ferguson, 1980; Boily, 1996; Hall et al., 1998 Boily, 1996

0.7

Schumacher et al., 1992

Pups (1–3 wk) Pups (4–10 wk)

3.3 3.5–4.3

Kirby, 1990 Kirby, 1990

Lactating Molting females

3.5 4.3

Kirby, 1990 Kirby, 1990 (Continued)


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TABLE 2 Reported Circulating Concentrations (µg/dl) of Thyroxine in Marine Mammals (continued) Species Mirounga leonina (southern elephant seal) Pagophilus groenlandicus (harp seal)

Specimens

Phoca vitulina (harbor seal)

Reference

Neonate

2.9

Little, 1991

Weaned

1.3

Little, 1991

Neonates

1.3–19.1

Pups (1000 IU/l) (Bossart, unpubl. data). Similar trends may be expected in pinnipeds, sea otters, and polar bears. In polar bears, GGT may fluctuate seasonally (Cattet, pers. comm.). Obstructive liver disease and cholestasis are uncommon in manatees; therefore, the significance of this enzyme in this species is unclear. GGT may be useful as a marker for passive immunoglobulin transfer in neonatal marine mammals. In domestic mammals, colostrum and milk have high GGT activity, with nursing animals having increased serum GGT activity. Meyer and Harvey (1998) proposed the concept of age-related serum GGT in neonatal dairy calves, with values exceeding 200, 100, 75, and 65 IU/l at 1, 4, 7, and 10 days of age, respectively.


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GGT liver disease (species-specific), including: primary acute obstructive liver disease cholestasis chronic obstructive liver disease (cirrhosis) skeletal muscle injury

Lipemia can have variable effects on GGT depending on methodology. GGT may be elevated in nursing immature animals; levels may slowly decline with age. Corticosteroids and phenobarbital can cause elevations of serum GGT.

Alkaline Phosphatase (ALP) ALP is an enzyme located on the cell membranes of a variety of tissues including liver, kidney, bone, heart, and skeletal muscle. In terrestrial mammals, ALP isoenzymes can be separated by electrophoresis, and two of these are diagnostically important: hepatobiliary and bone ALP. In terrestrial species, increased serum ALP levels are present in growing, young animals, and animals with bone and hepatobiliary disease. ALP shows minimal activity in normal hepatic tissue, but becomes markedly increased in serum, subsequent to increased enzyme production stimulated by impaired bile flow. Although detailed studies have not been conducted, similar trends for serum ALP activity may also exist in many marine mammal species. High serum ALP levels are seen in young, healthy, rapidly growing bottlenose dolphins, belugas, short-finned pilot whales, northern elephant seals, polar bears, and in some cases of bone and/or hepatobiliary disease in dolphins and pinnipeds (Lee et al., 1977; MacDonald, 1981). However, considerable species variation occurs in the diagnostic application of ALP to domestic animals, and similar variation also may exist in marine mammals. In marine mammals, ALP activity has been described as liver specific, with elevations in adult animals indicating liver damage (Medway and Geraci, 1986). However, in some dolphin species, infectious and noninfectious hepatopathies are typically not associated with serum elevations of ALP. Bottlenose dolphins and Pacific white-sided dolphins with histopathological confirmation of obstructive liver disease, including chronic fibrosing cholangiohepatitis, hepatic cirrhosis, chronic–active viral hepatitis, and hepatic hemochromatosis, did not have elevated serum ALP, but did have marked elevations of serum AST, ALT, and GGT (Bossart, 1984; Bossart et al., 1990). Furthermore, serum ALP activity >1000 IU/l was seen in an adult bottlenose dolphin with renal calculi, but with no clinicopathological evidence of hepatic disease (Dougherty, pers. comm.). In some dolphin species, serum ALP activity is a useful prognostic indicator (Fothergill et al., 1991). In critically ill, adult bottlenose dolphins and Pacific white-sided dolphins (with or without hepatic disease), serum ALP levels can typically and rapidly drop below 90 IU/l. If serum ALP levels remain decreased with clinical therapy, the prognosis for recovery is guarded. However, if serum ALP levels begin to return to normal, the prognosis is good. ALP levels can be used to evaluate nutritional status in cetaceans (Dover et al., 1993). The physiological mechanism to explain decreasing ALP levels with infectious disease is unknown. However, one possible explanation is that ALP functions in endotoxin detoxification, as observed in rats and humans (Poelstra et al., 1997). A similar mechanism may be present in some dolphins. Acute endotoxemia may consume and deplete available serum ALP, and so, with the resolution of endotoxemia, circulating levels of serum ALP return to normal. Another


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possible function may be related to food consumption; in rats and humans, serum ALP increased after consuming a meal high in lipid (Meyer and Harvey, 1998). Because inanition is typically a clinical component of disease in dolphins, this latter mechanism may account for rapid decreases of ALP levels observed in dolphins. Recently, serum ALP isoenzymes have been used as prognostic indicators for appendicular osteosarcoma in dogs (Ehrhart et al., 1998). The examination of serum ALP isoenzymes in dolphins may also provide useful prognostic information; ALP as a prognostic indicator in marine mammal species deserves further study. ALP liver disease (species-specific), including: obstructive hepatic disease/hepatic cirrhosis hepatitis bone disease chronic renal failure (decreased clearance) urolithasis (dolphins) young growing marine mammals (physiological) genetic abnormalities (not reported to date in marine mammals) geriatric marine mammals (physiological) hypothyroidism pernicious anemia decreased osteoblastic activity possible endotoxemia and inanition (some dolphin species)

Decreases in serum ALP can occur with hemolysis and with the use of the anticoagulants EDTA, oxalate, citrate, and fluoride. Lipemia and icterus can cause increases in serum ALP. Drugs that can increase ALP in some species include corticosteroids, anabolic steroids, sulfonamides, phenobarbital, phenytoin, primidone, azathioprine, barbiturates, and phenothiazines. Drugs that can decrease serum ALP include levamisole and theophylline.

Bilirubin Bilirubin is a yellow compound produced by the macrophage system from the degradation of heme from aged erythrocytes. Serum is typically yellow when bilirubin levels are greater than 1 mg/dl. Icterus may occur with accelerated destruction of erythrocytes, intrahepatic obstructive disease, or impairment of bile flow in the bile ducts (extrahepatic form). In the past, the use of the ratio of the unconjugated to conjugated bilirubin values was used to determine the etiopathogenesis of icterus. This was based primarily on extrapolation from human data. Recent studies in domestic animals indicate that species-specific variations in bilirubin metabolism preclude the reliable use of the ratio in the differentiation of the etiopathogenesis of icterus (Meyer and Harvey, 1998). In marine mammals, the kinetics of bilirubin metabolism are unknown. Therefore, use cautious interpretation of unconjugated and conjugated bilirubin serum concentrations. In general, elevated total serum bilirubin concentrations have been described with intrahepatic obstructive disease (cirrhosis or chronic–active cholangiohepatitis) in porpoises, Pacific white-sided dolphins, Atlantic bottlenose dolphins, and Risso’s dolphins (Grampus griseus) (Medway et al., 1966; Bossart, 1984; Bossart et al., 1990). In neonatal harbor seals, total bilirubin concentrations are high (>2 mg/dl), and similar to those reported in human neonates (Dierauf et al., 1984). Total serum bilirubin concentrations


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in healthy sea otters, polar bears, and manatees are similar to those in terrestrial mammals. In polar bears immobilized with phencyclidine, direct bilirubin values often decreased in animals that convulsed (Lee et al., 1977). Total Bilirubin liver disease (conjugated bilirubin elevation, but species-specific variations exist) bile flow impairment in the common bile duct (extrahepatic obstruction— conjugated bilirubin elevation, but species-specific variations exist) accelerated erythrocyte destruction (hemolysis, fetal hemoglobin eliminator— unconjugated bilirubin elevation, but species-specific variations exist)

Lipemia, hemolysis, ultraviolet light, and severe azotemia can decrease total bilirubin concentrations. Lipemia can cause a high serum total bilirubin concentration when measured by “wet” chemistry systems. Total bilirubin concentration can be elevated with the administration of acetaminophen, phenylbutazone, cephalosporins, sulfonamides, and propranolol.

Bile Acids Determination of serum bile acids has become a useful diagnostic test in domestic mammals and companion birds for detection of congenital portosystemic shunts, identification of chronic hepatitis/cirrhosis prior to the development of jaundice; it is also useful with therapy for monitoring the progression or resolution of hepatic disease. In some animals, fasting (FBA) and postprandial (PPBA) serum total bile acid concentrations are diagnostic indicators. The kinetics of the enterohepatic circulation of bile acids in marine mammals are not known; however, serum bile acid determination shows promise for diagnostic use in some marine mammal species. For example, in an Atlantic bottlenose dolphin with histologically confirmed hepatic hemochromatosis, with cirrhosis, random serum bile acid samples were >100 µmol/l (Bossart, unpubl. data). Further studies are needed to validate the use of this test in marine mammals. FBA and PPBA liver disease, and may result in: congenital portosystemic shunts chronic–active hepatitis cirrhosis, and/or monitoring therapy/prognosis

Lipemia and hemolysis can decrease serum total bile acid values.

Kidney-Associated Serum Analytes Urea Nitrogen and Creatinine Urea is found in the liver and represents the principal product of protein catabolism in most carnivorous and omnivorous species. Urea passes through the glomeruli and approximately 25 to 40% of filtered urea is reabsorbed in the renal tubules. Normal serum urea nitrogen (BUN) levels in most marine mammals are higher than in terrestrial mammals. Presumably, this is due to high dietary protein and fat. However, in polar bears, the BUN more closely parallels


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the BUN of terrestrial mammals (Lee et al., 1977). The common differentials for elevated BUN (due to prerenal causes) include dehydration, intestinal hemorrhage, septic shock, cardiac insufficiency, and postrelease catabolism/dehydration (speculated in manatees) (see Chapter 36, Nutrition). In contrast, liver failure and starvation cause BUN levels to decrease. Creatinine is formed in skeletal muscle metabolism and, as with BUN, serum creatinine is a crude index of glomerular filtration. However, creatinine is not reabsorbed by the renal tubules, and serum creatinine is not markedly influenced by diet or intestinal hemorrhage. Therefore, dehydration will initially cause a slightly elevated BUN with a serum creatinine concentration within the normal reference range. As dehydration, which causes decreased renal blood flow (prerenal azotemia), becomes more severe, both BUN and serum creatinine concentrations increase. Primary (renal) azotemia occurs with glomerular damage, and often results in a progressive proteinuria. Pharmacological agents known to produce azotemia in cetaceans are aminoglycosides (e.g., amikacin and gentamicin) and nonsteroidal anti-inflammatory agents (NSAIDs, e.g., flunixin and meglumine) (McBain and Reidarson, 1994). Post-renal azotemia occurs with urethral obstruction or is secondary to rupture of the urinary bladder. The same pre-renal and postrenal factors that affect BUN also influence creatinine. Therefore, the concurrent use of both analytes is useful for evaluating renal disease. One notable exception occurs in rehabilitated Florida manatees. Serum creatinine values at 2 to 4 weeks postrelease, ranged from 7 to 14 mg/dl (Lowe, Dougherty, Murphy, and Walsh, pers. comm.); BUN and other blood parameters remained within normal ranges. Clinically, these manatees appeared healthy, but exhibited mild to moderate weight loss compared with their prerelease weights. The cause of elevated creatinine is thought to be excessive skeletal muscle catabolism and nutritional factors (Manire et al., 1999). However, renal biopsies were not conducted to determine if glomerular damage or other lesions, which may influence creatinine levels, were present. BUN and Creatinine prerenal azotemia, including: dehydration cardiac insufficiency shock (septic or traumatic) gastrointestinal hemorrhage (BUN increase only) high protein diet (BUN only) post-release catabolism/dehydration (speculated in manatees) renal azotemia post-renal azotemia, including: obstruction of lower urinary tract and/or rupture of lower urinary tract liver failure (BUN only) starvation (BUN only)

Creatinine concentrations are artifactually increased with lipemia and ketonemia, and decreased with hemolysis, icterus, and when using the anticoagulants EDTA and oxalate. BUN concentration is increased when using the anticoagulants ammonium heparin and ammonium oxalate, and decreased when using sodium citrate and fluoride. Immature animals may have a lower BUN than adults. BUN and creatinine concentrations may increase with corticosteroids (BUN only), NSAIDs, aspirin, ibuprofen, aminoglycosides, cephalosporins (creatinine only), amphotericin B, ascorbic acid (creatinine only), cisplatin, furosemide (BUN only), salicylates, and some radiographic contrast media.


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Serum Proteins Hematocrit and Total Plasma Protein Important clinical information can be acquired by the simultaneous interpretation of the HCT and total plasma protein (TPP) concentrations (from microhematocrit capillary tube plasma refractometer determination). Various combinations of low, normal, or high HCT values and TPP values may indicate specific clinical problems. Comparing the two pieces of clinical information provides an efficient and simple method for gaining diagnostic information and is especially applicable to marine mammal field studies. The box shows diseases that may be diagnosed by examining changes in HCT and total plasma concentrations.

Interpretation of HCT and TPP Concentrations normal HCT with: high TPP = = normal TPP = low TPP = = = high HCT with: high TPP = normal TPP = = low TPP = low HCT with: high TPP = = = normal TPP = = = low TPP = =

increased globulin synthesis dehydration-masked anemia normal gastrointestinal protein loss renal protein loss severe liver disease dehydration dehydration-masked hypoproteinemia primary or secondary erythrocytosis protein loss with splenic contraction (not in cetaceans) chronic disease-associated anemia (common in marine mammals) lymphoproliferative diseases (rare) immunoblastic malignant lymphoma/multiple myeloma (rare) chronic blood loss hemolytic disease decreased RBC production acute or ongoing serious blood loss overhydration

Total Protein dehydration shock hypoalbuminemia malnutrition proteinuria protein-losing enteropathy

With lipemia, hemolysis, hyperglycemia, and severe azotemia, total protein can be increased using refractometer methodology. Icterus and sample dehydration can also increase total protein. The administration of corticosteroids and anabolic steroids can also increase total protein.


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Albumins and Globulins The interpretation of the kinetics of total protein, albumin, and globulin fractions (determined by electrophoresis) is receiving increased attention in the study of disease mechanisms in marine mammals. Albumin production is solely dependent on liver function and adequate nutrition. Albumin values and, consequently, albumin/globulin ratios are somewhat higher in cetaceans compared with domestic mammals. Automated serum chemistry analyzers using human standards typically give erroneous results with cetacean serum (Medway and Geraci, 1986). Accurate albumin values and clinically useful globulin fractions are therefore best obtained by serum protein electrophoresis. Marine mammals, like terrestrial mammals, appear to have a tremendous reserve capacity for hepatic albumin production. This limits the use of albumin as an early indicator of hepatic disease. Elevated levels of albumin occur with dehydration and shock, producing relative hyperalbuminemias. Decreasing albumin levels are observed with malnutrition, protein-losing nephropathies, gastrointestinal disease (especially with parasitism, maldigestion/malassimilation syndromes, and protein-losing enteropathies), advanced hepatic disease, downregulation of albumin production secondary to hyperglobulinemia, hemorrhage, and severe and extensive skin lesions with epidermal compromise. Albumin dehydration shock malnutrition renal disease (protein-losing nephropathy) gastrointestinal disease, including: parasites maldigestion protein-losing enteropathy advanced hepatic disease downregulation of albumin production secondary to hyperglobulinemia hemorrhage (internal body cavity or external hemorrhage, including gastrointestinal) severe and extensive skin lesions with epidermal compromise (e.g., burns) Note: Absolute hyperalbuminemia is nonexistent; increases are relative.

Hemolysis increases albumin concentration. Lipemia decreases albumin concentration, as can estrogen administration. Globulin concentrations are determined by subtracting total serum albumin from serum protein. Individual globulin fractions are separated and quantified by serum protein electrophoresis (SPEP), which separates serum proteins based on charge densities and resultant mobility in an electric field. The absolute value for each protein fraction is calculated by multiplying the percentage of each fraction by the chemically determined total serum protein concentration. Protein fractions include prealbumin (seen in some cetaceans and neonatal marine mammals), albumin, α-globulins (α-1 and α-2), β-globulins (β-1 and β-2) and γ-globulins. SPEP can provide clinically useful information and is the preferred method for determining albumin concentration in marine mammals. Interestingly, globulin levels were significantly greater in free-ranging belugas and bottlenose dolphins compared with captive individuals (see Table 1). The significance of these findings is unknown at this time. To better define these differences, SPEP and immune function tests may be in order.


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α-Globulins are acute-phase proteins that include haptoglobin, lipoproteins, and antitrypsin. β-Globulins include complement, hemopexin, transferrin, and fibrinogen. Acute-phase proteins are important early markers of acute inflammatory disease in some marine mammal species (see Chapter 12, Immunology). These acute-phase proteins may be elevated in inflammatory or infectious disease prior to other clinicopathological signs appearing. Additionally, some immunoglobulins (IgM and IgE) may also migrate in the β range. Immunoglobulins or antibodies are glycoproteins produced by plasma cells that proliferate in response to antigenic stimulation of B-lymphocytes (see Chapter 12, Immunology). Seldom can a specific diagnosis be made with SPEP alone. However, dysproteinemias can be associated with some types of disease processes. SPEP does provide a rationale for further diagnostic studies, particularly with dysproteinemias. For example, the absence of γ-globulins in precolostral or colostrum-deprived neonatal cetaceans, harbor seals, and manatees can be readily demonstrated by SPEP (McBain and Reidarson, 1998). In dolphins, acquired immunodeficiency can be seen with chronic disease and suspected immunological compromise (Bossart, 1984). Additionally, β-γ bridging can be seen in some cases of dolphin hepatitis (Bossart et al., 1990). This bridging has also been seen with hepatitis in domestic animals (Kaneko, 1980). To characterize dysproteinemias further, the disease states in the box below relate to changes in individual globulin fractions. Globulins (by SPEP) α-globulins with: acute inflammatory disease (can occur before other diagnostic signs of inflammation occur) severe active hepatitis acute glomerular disease and the nephrotic syndrome β-globulins with: acute hepatitis suppurative dermatopathies nephrotic syndrome β-globulins and β–γ bridging with: chronic–active hepatitis in bottlenose dolphins (suspected bacterial etiology) hepadnavirus chronic–active hepatitis in Pacific white-sided dolphins hepatic hemochromatosis in Atlantic bottlenose dolphins γ-globulins—broadband (polyclonal) increases with: chronic inflammatory disease (usually associated with a concomitant decrease in albumin as a result of decreased synthesis; observed in cetaceans and manatees (Ridgway, 1972; Bossart and Bigger, 1994) chronic hepatitis, pulmonary/hepatic abscessation, other suppurative disease processes (the polyclonal increase with suppurative disease is usually more marked, and the hypoalbuminemia more severe than in chronic inflammatory disease; this likely reflects more intense antigenic stimulation), neoplasia γ-globulins—sharp band (monoclonal) increases with: neoplasia (suspected in delphinid immunoblastic malignant lymphoma; Bossart et al., 1997) non-neoplastic plasma cell proliferation (plasmacytic enterocolitis, idiopathic) γ-globulins with: fetal serum precolostral neonate acquired immunodeficiency of chronic-disease states (suggesting humoral immunological exhaustion)

Immature marine mammals, when compared with adults, generally have lower globulin concentrations. Hemolysis and lipemia can cause erroneous SPEP protein values.


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Electrolytes Serum electrolytes are useful aids in helping assess hydration status and some gastrointestinal and endocrine conditions in marine mammals. In general, electrolyte values in marine mammals are similar to those reported for other marine and terrestrial mammals (Geraci, 1971; Greenwood et al., 1971; Lane et al., 1972; Ridgway, 1972; White et al., 1976; Englehardt, 1979).

Sodium Pinniped hyponatremia is a disorder principally of phocid seals and is characterized by a sudden or gradual decrease in circulating levels of sodium (Geraci, 1972). Normally, serum sodium ranges from 120 to 147 mEq/l (see Chapter 30, Intensive Care; Chapter 36, Nutrition; Chapter 41, Seals and Sea Lions). Other causes of hyponatremia include gastrointestinal loss due to vomiting or diarrhea, congestive heart failure with edema, hypoadrenocorticism, diuretic treatment, diabetes mellitus, and chylothorax. Causes of hypernatremia include dehydration secondary to inadequate water intake or excessive water loss, increased salt intake or intravenous administration, hepatic cirrhosis, renal failure, diabetes insipidus, hyperaldosteronism, and diuretic treatment. Sodium dehydration secondary to inadequate water intake or excessive water loss hepatic cirrhosis renal failure diabetes insipidus hyperaldosteronism diuretic treatment increased salt intake or intravenous administration gastrointestinal loss (vomiting, diarrhea) congestive heart failure with edema hypoadrenocorticism diuretic treatment diabetes mellitus chylothorax

Sodium concentrations may be decreased when a lipemic sample is measured by flame photometry or indirect potentiometry. Hyperglobulinemia may also decrease sodium levels. The sodium salts of the anticoagulants EDTA, fluoride, and heparin can increase sodium concentrations. Additionally, sample dehydration can increase sodium levels. Drugs that can increase sodium concentrations include corticosteroids, mineralocorticoids, phenylbutazone, androgens, and sodium bicarbonate. Drugs that can decrease sodium concentrations include furosemide and some NSAIDs.

Potassium Potassium is sometimes elevated following severe physical exertion in seals and dolphins (Medway and Geraci, 1978). Other causes of hyperkalemia include renal failure, urethral obstruction, hypoadrenocorticism, acidosis, rhabdomyolysis (in some stranded cetaceans; Bossart and Trimm, 1993), and diffuse cellular necrosis secondary to shock. Causes of hypokalemia include gastrointestinal loss due to vomiting and/or diarrhea, diuretics, hyperaldosteronism, and reduced dietary intake.


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Potassium renal failure including urethral obstruction hypoadrenocorticism acidosis rhabdomyolysis (in some stranded cetaceans) diffuse cellular necrosis secondary to shock gastrointestinal loss (e.g., vomiting, diarrhea) diuretics hyperaldosteronism reduced dietary intake

Increases (often marked) in potassium concentrations are not uncommon in marine mammals samples that are hemolyzed and in animals with excessive leukocytosis or thrombocytosis (due to cell leakage during clotting). By using plasma samples for potassium measurements, the latter artifact can be avoided. Increases can also occur with the anticoagulants EDTA, citrate, and fluoride, which contain potassium salts. Decreases in potassium can occur with lipemia (using flame photometric methods), hyperglycemia, and severe azotemia (using dry reagent methods). Drugs that can increase potassium concentrations include NSAIDs, androgens, heparin, and propranolol. Drugs that can decrease potassium concentrations are corticosteroids, mineralocorticoids, aspirin, insulin, amphotericin B, flucytosine, furosemide, and sodium bicarbonate.

Chloride Chloride levels can be elevated due to increased salt or saltwater intake, dehydration, and renal tubular acidosis. They decline with protracted vomiting, diarrhea, and metabolic acidosis. Chloride dehydration renal tubular acidosis metabolic acidosis prolonged vomiting

Chloride can increase artifactually with icterus. Decreases in chloride concentrations can occur with lipemia and hyperproteinemia (with flame photometric methods).

Total Carbon Dioxide Total CO2 is not a direct determination of bicarbonate concentration; total CO2 represents dissolved CO2 plus bicarbonate plus carbonic acid. However, the determination of total CO2 may be of value in characterizing disturbances in acid–base balance. −

Total CO 2 = Dissolved CO 2 + HCO 3 + H 2 CO 3

(2)

Bicarbonate concentration can be estimated in terrestrial mammals by subtracting 1.2 from the total CO2 value (Coles, 1986).


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Total CO2 metabolic alkalosis partially compensated respiratory acidosis metabolic acidosis partially compensated respiratory alkalosis

Calcium, Phosphorus, and Magnesium Calcium, phosphorus, and magnesium homeostasis are similar, in that the same conditions and hormones tend to regulate their excretions. Calcium

Plasma calcium exists in an ionized (or free) form (approximately 50%), bound to albumin (approximately 45%), and as anions (approximately 5%). The ionized form is the physiologically active form, and the pH of extracellular fluids and total protein concentrations can change plasma levels. Acidosis can cause an increase in ionized calcium; alkalosis can cause a decrease in ionized calcium. An increase in total protein concentration results in an increase in the calcium level. Reduced total protein concentration has the opposite effect. Calcium hyperalbuminemia (dehydration) primary hyperparathyroidism hypoadrenocorticism hypervitaminosis D renal disease hypercalcemia of neoplasia (pseudohyperparathyroidism) osteolytic bone lesions plant toxicity (species-specific) calciferol rodenticides some granulomatous diseases hypoalbuminemia alkalosis hypoparathyroidism secondary renal hyperparathyroidism necrotizing pancreatitis nutritional imbalances (hypovitaminosis D, excess phosphorus) eclampsia or parturient paresis hypomagnesemic tetany intestinal malabsorption hypercalcitoninism transport tetany

Increases in calcium concentrations can occur with lipemia and hemolysis. Concentrations of calcium decrease with the anticoagulants EDTA, oxalate, and citrate. Drugs that can increase calcium concentrations are anabolic steroids, androgens, estrogens, and progesterone. Decreases can occur with corticosteroids, anticonvulsants, and bicarbonate treatment for salicylate toxicity.


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Phosphorus

Phosphorus decreased glomerular filtration rate (pre-renal, renal, or postrenal azotemia) dietary phosphorus excess hypervitaminosis D osteolytic bone disease massive cell lysis (rhabdomyolysis associated with stranding; Bossart and Trimm, 1993) hypoparathyroidism with normal glomerular filtration hypercalcemia of malignancy with normal glomerular filtration ethylene glycol toxicity (sudden rise) primary hyperparathyroidism dietary calcium deficiency hypovitaminosis D hyperadrenocorticism eclampsia/parturient paresis starvation/malabsorption vitamin D intoxication enteral alimentation diabetes mellitus and ketoacidosis chronic renal failure

Increases in phosphorus concentrations occur with lipemia, hemolysis, and icterus. Blood samples stored too long before analysis can show increases in phosphorus due to its release from erythrocytes. Immature growing animals have phosphorus levels above adult normals. Drugs including anticonvulsants, insulin, phenothiazines, and salicylates can lower phosphorus concentrations. Magnesium

Severe hypomagnesemia impairs parathyroid hormone (PTH) secretion, and magnesium supplementation may be required to restore normal PTH and calcium concentrations. In most mammals, the plasma magnesium concentration correlates poorly with the total body status (Elin, 1994). An accurate assessment of magnesium necessitates complex metabolic studies. Potassium may serve as a surrogate marker for magnesium because both are present as intracellular cations. Magnesium levels in marine mammals are not commonly reported. Magnesium ↓

critical care marine mammal patients, and may lead to cardiac dysfunction

Increases in magnesium concentrations can occur with hemolysis. Magnesium decreases with icterus and EDTA anticoagulant. Progesterone can increase magnesium concentrations. Drugs that can decrease magnesium concentrations include aminoglycosides, amphotericin B, furosemide, and cisplatin.


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Miscellaneous Serum Analytes Uric Acid Uric acid levels are unusually high in young phocids (2 to 5 mg/dl) and freshwater dolphins (10 to 12 mg/dl) (Medway and Geraci, 1986). The measurement of uric acid is of doubtful value for assessing liver and renal disease in marine mammals. Uric acid concentrations may be elevated in dehydrated seals, dolphins, and manatees. Uric Acid dehydration gout

Creatinine Phosphokinase Creatinine phosphokinase (CK, CPK) is found as three tissue-specific isoenzymes in skeletal muscle, myocardium, and brain of terrestrial mammals. It appears that in marine mammals most increases in plasma CPK occur with skeletal muscle injury associated with strenuous activity, transport, surgery, stranding, seizures, and intramuscular injections. In sea otters, CPK concentrations have a wide range, possibly as a result of bleeding procedures, struggling, and/or capture (Williams and Pulley, 1983). The use of CPK isoenzymes has not been widely investigated in marine mammals. Creatinine Phosphokinase (CK, CPK) skeletal muscle disease myocardial disease central nervous system disease handling stress/stranding

Increases in CPK concentrations can occur with hemolysis. Decreases can occur with the anticoagulants EDTA, oxalate, and citrate. Drugs that can increase plasma CPK concentrations include corticosteroids.

Hemostatic Parameters Blood Types Specific blood types have been described in whales, dolphins, and seals (Ridgway, 1972; Bonner and Fogden, 1974; Cornell et al., 1981).

Screening for Hemostatic Disorders A battery of hemostatic tests are usually required to diagnose hemostatic disorders. These tests include platelet count, mean platelet volume (MPV), bleeding time, activated clotting time, activated partial thromboplastin time (APTT), prothrombin time (PT), thrombin clotting time (TCT), fibrinogen, and fibrin degradation products. Most of these tests require blood collected in citrate tubes (blue top) with a proper blood to anticoagulant ratio (9 : 1). Tests are time and temperature dependent. Tests must be performed


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within 2 hours at room temperature or 12 hours if samples are refrigerated. If refrigeration is impossible, a less acceptable, but functional method can be used: samples are centrifuged and plasma separated for later testing. Always collect and run a control sample from a healthy animal of the same species along with the test sample. Coagulation dynamics of most marine mammal species has only been minimally investigated. Odontocetes are known to have deficiencies in Hageman factor (Factor XII) activity and Fletcher Factor activity (plasma prekallikrein) (Lewis et al., 1969; Robinson et al., 1969). Additionally, sei whales (Balaenoptera borealis) had prolonged partial thromboplastin time with no detectable Factor XII, XI, or Fletcher Factor (Saito et al., 1976). The functional significance of these findings is unclear because these marine mammals did not appear to have hemostatic disorders. Coagulation assay parameters have been established in northern elephants seals because of the frequency of disseminated intravascular coagulation in this species (Gulland et al., 1996). Some electronic cell counters cannot accurately count platelets in whole blood samples from marine mammals; the presence of platelet clumps in many marine mammal blood samples often results in erroneously low platelet counts. Stained peripheral blood smears must be examined to verify low platelet counts. Bleeding time is a crude, but simple, hemostatic test that generally involves penetration of oral mucous membranes with a lancet. Left undisturbed, bleeding will usually stop in less than 5 min. Often the first sign of a hemostatic disorder is prolonged bleeding following venipuncture. If bleeding time is prolonged, but the platelet count is normal, coagulation tests are recommended. Because minimal data exist on clotting mechanisms in marine mammals, the clinician would be wise to refer to the domestic animal literature for possible data extrapolation. Hemostatic disorders appear to be uncommon in marine mammals with three notable exceptions. Liver disease in dolphins has been associated with increased clotting time, prolonged PT, prolonged APTT, and decreased fibrinogen (Bossart et al., 1990). Autoimmune hemolytic anemia and thrombocytopenia were suspected in a bottlenose dolphin, based on macroscopic agglutination and a positive dolphin-adapted Coombs test (Patterson, pers. comm.). Widespread microvascular thrombosis and hemorrhage suggestive of disseminated intravascular coagulation (DIC) have been observed in some marine mammals at post-mortem. DIC is a recognized abnormality of hemostatic function that develops secondarily to an underlying or primary pathological process (Slappendal, 1988). This thrombohemorrhagic disorder paradoxically results in simultaneous widespread microvascular thrombosis and hemorrhage. The paradox results from the simultaneous and excessive, or unbalanced, generation of thrombin and plasmin (Bateman et al., 1999). Systemic circulation of thrombin causes widespread microvascular thrombosis resulting in tissue hypoxia, acidosis, cell death, and organ failure. The consumption of coagulation factors and platelets in the formation of these thrombi creates a tendency for hemorrhage. Systemic circulation of plasmin results in widespread lysis of clotting factors, which contributes to further hemorrhage. In terrestrial mammals, conditions that may result in DIC include bacterial septicemia, viremia, protozoal parasites, liver disease, and traumatic shock. Microscopic lesions suggestive of acute, or decompensated, DIC accompanied by shock have been seen in manatees following massive boat-collision trauma, in a bottlenose dolphin with suspected acute morbillivirus infection (Bossart, unpubl. data), and in northern elephant seals with bacterial and parasitic infections (Gulland et al., 1996; 1997). Additionally, a form of consumptive coagulopathy has been postulated in manatees with nonlethal, but clinical, neurotoxic brevetoxicosis (Murphy, pers. comm.).

Prothrombin Time and Partial Prothrombin Time PT in humans can be used as a prognostic indicator for liver disease, specifically viral hepatopathies. In some dolphin species with liver disease, PT has been used as a prognostic indicator.


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One Pacific white-sided dolphin and three bottlenose dolphins had acute hepatopathies characterized by marked elevations of ALT, AST, and GGT, with prolonged PTs. In the white-sided dolphin, PT was approximately 22 s (control = 11 s). In three bottlenose dolphins, PTs ranged up to 30 s. Clinicians need to consider a guarded prognosis in a dolphin with a persistently elevated PT. Alternatively, an improving prognosis exists with normalization of PT and a subsequent drop in hepatic-specific enzymes (Bossart et al., 1990). Interestingly, partial prothrombin times are normally elevated (>2 min) in bottlenose dolphins, pilot whales, and killer whales (Reidarson, unpubl. data).

Markers of Inflammation A detailed account of clinical markers of inflammation is given in Chapter 12, Immunology.

Erythrocyte Sedimentation Rate Erythrocyte sedimentation rate (ESR) is a measurement (in mm) of the distance erythrocytes fall through a vertical suspension of anticoagulant over time. ESR is a nonspecific test that can be useful in monitoring the presence and intensity of inflammation in most dolphin species (Schroeder, pers. comm.). In dolphins, the magnitude of inflammation is directly related to the rate with which RBCs fall in a standard vertically positioned tube (i.e., ESR increases with increasing inflammatory disease). Most increases in ESR are also associated with increases in plasma fibrinogen; thus, quantitative determination of this acute inflammatory phase protein has almost completely replaced ESR as a laboratory diagnostic test, at least in domestic animals and humans. In bottlenose dolphins, normal ESR ranges from 1 to 56 mm and falls in 60 min (mean = 11 mm, n = 210; Schroeder, unpubl. data). With resolution of the inflammatory response and/or tissue injury, ESR declines to within normal ranges. Consequently, ESR can still be used in dolphins as a prognostic indicator. Artificially low ESR values can be seen in dehydrated individuals, due to hyperviscous serum (Reidarson, unpubl. data).

Serum Iron The majority of iron found in mammals is bound to hemoglobin, myoglobin, and cytochrome proteins. The remaining iron is either bound by other iron-binding proteins (transferrin, lactoferrin, and ferritin) or exists in small amounts in a free form. In domestic animals, changes in serum iron have been used as indicators of inflammation. In the acutephase inflammatory response, iron is sequestered by iron-binding proteins, making it unavailable for invading pathogens, and thus decreasing the chance for infection. Acutephase proteins help mediate this iron sequestration. Similar decreases in serum iron have been described for dolphins with inflammatory conditions involving trauma, parasitism, infectious disease, and metabolic derangements (Medway and Geraci, 1986; Fenwick et al., 1988; McBain, 1996). Furthermore, these studies demonstrated that trends in serum iron levels were important when evaluating clinical condition and prognosis in marine mammals (i.e., the elevation of serum iron to normal concentrations following a clinical illness is generally a favorable prognostic indicator). In one study, normal ranges of iron in dolphins were serum iron = 116 to 320 µg/dl, unbound iron-binding capacity (UIBC) = 100 to 564 µg/dl, and iron saturation = 17 to 65% (Medway and Geraci, 1986). Elevated serum iron levels have been observed in two Atlantic bottlenose dolphins with hemochromatosis confirmed at necropsy (Bossart, unpubl. data). Other causes of hyperferremia are trauma and overzealous iron supplementation.


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Serum Iron hemolysis trauma hemochromatosis (dolphins) overzealous iron supplementation iron deficiencies active erythrogenesis infectious disease (especially bacterial disease) physiological stress

Increases in serum iron occur with hemolysis. Decreases occur with the anticoagulants EDTA, oxalate, and fluoride. Immature animals may have lower levels of serum iron than adults.

Bone Marrow Evaluation Bone marrow aspirates and biopsies can be obtained from marine mammals from the central vertebral bodies of the tail stock in cetaceans and manatees, and from the pelvis or femoral trochanter in pinnipeds, sea otters, and polar bears. Radiographic localization of the radiolucent vertebral bone marrow space is often useful prior to biopsy in cetaceans (Walsh, pers. comm.). The technique for aspiration or core biopsy is by standard procedures used for small animals (Cotter and Blue, 1985). Indications for bone marrow evaluation include nonregenerative anemia, aplastic anemia, and abnormalities in circulating numbers or types of leukocytes or platelets. A peripheral blood smear should accompany a bone marrow aspirate and/or core biopsy. Additionally, core biopsies are recommended over aspirates for anemic animals.

Urinalysis Minimal data exist about urine output, collection, and characteristics in marine mammals. Urine can be collected by free-catch (voided), catheterization, and/or cystocentesis (see Chapters 40 through 45). Catheterization of manatees can be difficult because the penis cannot be extruded manually, and the female’s urogenital slit is difficult to retract. An otoscope with a large-core speculum may be useful. Firm and constant external digital pressure over the urinary bladder may induce urination in this species, and ultrasound-guided cystocentesis is also a possible collection technique. Urinalysis is best performed on freshly collected urine, although refrigerated specimens are generally acceptable for as long as 6 hours. Physical and chemical properties to examine include appearance, specific gravity, pH, protein, glucose, ketones, occult blood, bilirubin, and microscopic sediment (leukocytes, erythrocytes, epithelial cells, casts, bacteria, yeast, fungi, sperm, and crystals). Enzymes, including GGT and ALP, can also be measured. In domestic animals these are markers for renal tubular injury. Urine electrolytes have been measured in some marine mammals with confusing results (Bossart, unpubl. data). Results can be confusing because of the absence of control marine mammal population reference values and the numerous variables that can abruptly affect the appearance of electrolytes in urine. A diet high in fish protein fed to cetaceans, pinnipeds, sea otters, or polar bears results in deep amber-colored transparent urine with a pH of approximately 6 (Medway and Geraci, 1986). Manatees normally have amber-colored transparent urine with a pH of 6 to 7.5.


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Conclusion It is clear from these discussions and case studies (to follow), that in the last decade, knowledge of marine mammal clinical laboratory medicine has greatly expanded, not only offering the clinician additional diagnostic tests to utilize, but also providing sizable areas within clinical laboratory medicine in which to conduct bench science and research. Following are a few case studies demonstrating the use of current diagnostic testing in marine mammals.

Clinical Cases Cetaceans CASE 1 (Miller et al., 1999)—Bottlenose Dolphin History

A clinically normal female primiparous 15-year-old bottlenose dolphin became anorexic and soon thereafter delivered a dead female fetus without complications. Clinicopathological Findings

The cow had a moderately elevated ESR (30 min @ 1 hour), mildly elevated inorganic phosphate (5.2 mg/dl), and mildly elevated serum creatinine (2.4 mg/dl). All other hematological and serum analytes were normal. A Brucella sp. was isolated from the placenta that had biochemical and oxidative metabolic profiles, which resembled, but did not match, the profiles in established species and biovars of Brucella. A severe multifocal suppurative placentitis was present with a necrotizing vasculitis. Brucella antigens were detected with immunohistochemistry. The cow became pregnant again soon after the abortion and subsequently produced and raised a healthy calf. Discussion

Brucella-induced abortions and infections have been described in three bottlenose dolphins (Chapter 16, Bacterial Diseases). Microbiology, specific polymerase chain reaction, and pulsedgel electrophoresis results supported the designation of an additional genomic group, B. delphini, for isolates adapted to dolphins. Current serological tests that are reliable for diagnosing known Brucella species are unreliable in detecting dolphin brucellosis. This disease likely occurs naturally and can adversely impact reproduction in dolphins. The zoonotic significance is unknown. CASE 2 (Reidarson et al., 1998)—Bottlenose Dolphin History

A 5-year-old bottlenose dolphin developed a harsh cough. The dolphin was slightly underweight and had a prolonged inspiration and expiration. Clinicopathological Findings

Hematological findings indicated nonspecific inflammatory disease. Thoracic radiographs illustrated a 4 to 5 cm focal alveolar pattern in the left caudal lung lobe with associated interstitial changes. Blowhole cytological studies were unremarkable and a mixed bacterial blowhole culture grew Morganella morganii, Staphylococcus intermedius, and Vibrio alginolyticus. Treatment

The dolphin was initially treated for what was believed to be a bacterial pneumonia with cefuroxime at 20 mg/kg BID PO. Progress

The hematological data improved, but the cough persisted.


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Additional Clinicopathological Findings

Additional blowhole cultures yielded Pseudomonas aeruginosa, Streptococcus pyogenes, and Staphylococcus epidermidis. Two antibody bands specific to Aspergillus fumigatus were identified using serological immunodiffusion. Retrospective sampling revealed a single band associated with the first clinical signs and no bands 3 months earlier. Further Treatment

Itraconazole 5 mg/kg BID PO and amoxicillin/clavulanic acid at 5 mg/kg BID PO were given. Because of worsening cough, bronchoscopy was performed, bronchoalveolar lavages were performed, and a brush biopsy of a 1-cm raised yellow lesion of the left side mainstem bronchus was conducted. Lavage cytological findings demonstrated branching septate hyphae admixed with neutrophils; the hyphae were identified as A. fumigatus. Itraconazole therapy was continued for 9 months. Bronchoscopic examination 2 months after the first bronchoscopy demonstrated bronchial lesion resolution and the absence of fungi from lavage fluid, with improvement of cough and normal body weight. CASE 3 (Stetter et al., 1999)—Bottlenose Dolphin History

An adult, male bottlenose dolphin with no clinical signs of disease had elevated blood lead concentrations (92 µg/dl). Clinicopathological Findings

Radiographs illustrated the presence of metallic objects in the dolphin’s first stomach compartment. All other blood parameters were normal. Treatment

Chelation therapy was initiated using dimercaptosuccinic acid, 600 mg BID PO for 5 to 7 days, followed by several weeks without therapy for a total of nine treatment cycles. Other oral treatment included mineral oil, sucralfate, nystatin, and enrofloxacin. Endoscopy and gastric lavage were used to remove the lead material from the first stomach compartment. Discussion

Several blood samples over an 8-month period demonstrated a steady decline in lead concentrations. The dolphin remained clinically normal with blood lead concentrations of less than 10 µg/dl over the next year. CASE 4 (Bossart and Eimstad, 1988)—Killer Whale History

An adult female, in captivity since the early 1970s, that had been vaccinated for erysipelas annually from 1971 to 1977, developed a focal progressive vesicular glossitis. Through a trained behavior, the whale allowed the tongue vesicle to be aspirated aseptically. Cytologically, the vesicle contained degenerating toxic neutrophils and hemorrhage. A bacterial culture of the lesion grew a pure isolate of Erysipelothrix rhusiopathiae. Diagnosis

Erysipelothrix vesicular glossitis. Treatment

The Erysipelothrix was sensitive to a wide range of antibiotics. During initial examination, the whale was treated with cefadroxil (20 g BID PO), and sensitivity results indicated the bacteria were responsive to this antibiotic.


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Discussion

This was an unusual presentation for erysipelas in a delphinid. The literature describes erysipelas as either a peracute septicemic disease in dolphins and pigs or a treatable dermatological disease characterized by rhomboid plaques of the skin. The vesicular glossitis was probably introduced by a fish spine. However, on six separate occasions, Erysipelothrix was not cultured from fish or pool water. As the tongue lesion progressively enlarged, it eventually ruptured, forming an ulcer approximately 20 × 10 × 5 cm. The ulcer granulated in approximately 4 weeks. In reported cases of cutaneous delphinid erysipelas, a leukocytosis or change in behavior denoted illness. In this whale, there was no leukocytosis or change in behavior, and appetite was normal. There were no hematological abnormalities, and a detailed data bank existed for the animal. It is unknown whether the animal’s past vaccinations with a killed bacteria had caused this atypical morphological expression. The animal at the time of vesicle aspiration had a positive erysipelas titer of 1 : 1045, suggesting either infectious exposure or prior immunization. CASE 5 (Bossart et al., 1996)—Killer Whale History

An adult, male killer whale in captivity developed bilateral axillary and peduncular tail stalk lesions, which had a 10-year cyclical pattern of proliferation and regression. The proliferative nature of the lesions was often extensive, extending from the axillae to the lateral thorax and involving up to approximately 40 × 30 cm areas. Clinicopathological Findings

Histological and electron microscopic evaluation of skin lesions were consistent with cutaneous papillomatosis due to a papillomavirus. Routine hematological and serum chemical parameters were generally within normal ranges for killer whales. Specialized immunohematological studies supported the hypothesis of an underlying immunological dysfunction. Discussion

The clinical presentation of these lesions was unusual compared with other mammalian species with cutaneous viral papillomatosis. The bilateral symmetry of the lesions was unique, as most terrestrial species with this disease have focal or generalized and randomly distributed lesions. The erratic cyclical pattern of partial lesion resolution and proliferation was also unusual. In most mammalian species, cutaneous viral papillomatosis is self-limiting and spontaneously regressive. The cyclic nature of this disease indicated a possible immunological component in the disease pathogenesis. CASE 6 (Bossart et al., 1990)—Pacific White-Sided Dolphin History

An alteration in training performance and behavior in an adult female approximately 22 years of age, in captivity since the early 1970s, was noted and blood was collected. Clinicopathological Findings (Day 1)

There were mild to moderate elevations of AST, ALT, and GGT, and a mild leukocytosis, with relative neutrophilia, relative lymphopenia, and relative eosinopenia. There was also a regenerative left shift. Treatment

Because the animal’s appetite remained normal, 3 g of cephalexin BID PO was administered for 21 days and the animal was allowed to rest. A week later, icterus at the site of an old erysipelas scar, as well as the sclera, was noted.


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Subsequent Clinicopathological Findings (Day 9 to 44)

There was a marked increase in ALT (4165 IU/l) (normal ALT for this animal was between 53 and 64 IU/l), a marked increase in AST (>2500 IU/l) (normal for this animal was ∼190 IU/l), and an elevation in total bilirubin (0.9 mg/dl) (indirect and direct bilirubin values approximately the same). There was marked elevation in GGT (>1000 IU/l) (normal for this animal 3 was ∼40 IU/l), and WBC count fluctuated between 15,800 and 20,400/mm (normal range for 3 this animal was between 4000 and 6000/mm ), and mild to moderate hypergammaglobulinemia (3.1 mg/dl). AP remained within the normal range for this animal. Elevated PT (∼22 s, with a control time of 11 s). Hypoglycemia (48 mg/dl) (normal range = 92 to 113 mg/dl). During the second week of infection, a serological survey for human viral hepatitis was done for this animal. Results of the survey were as follows: negative results for hepatitis A antibody (anti-HA), hepatitis B surface antigen (HBsAg), and hepatitis B antigen (HBeAg). Positive results for hepatitis B core antibody (anti-HBc), hepatitis B surface antibody (anti-HBs), and hepatitis B viral DNA (HBv). Additional Treatment

Penicillin G was administered at 400,000 IU BID PO for 14 days. Menadiol sodium phosphate (5 mg BID PO), oral dextrose, and additional multivitamin supplementation were also given. Diagnosis

The diagnosis was hepadnavirus hepatitis. In humans, anti-HBc is the most specific test for hepatitis B infection. HBv in humans indicates active viral replication. The clinicopathological signs observed for this dolphin were consistent with a resolving hepatitis B-like infection. Discussion

It is not known how this disease was transmitted. Hepatitis B titers were examined for the animal’s trainers and veterinarian. All serological tests on humans were negative for viral hepatopathies. As the disease progressed, the dolphin’s BUN decreased from 53 to 26 mg/dl. This can be associated with hepatic failure in other species. A decrease in glucose was also noted, consistent with hepatic failure because there was reduced hepatic gluconeogenesis. The polyclonal hypergammaglobulinemia suggested an immunological response to the disease. The elevated GGT was probably due to obstructive hepatic disease. The PT returned to normal in this animal, but, periodically, cyclic episodes of active hepatopathy were observed. A killer whale housed with this dolphin had anti-HBs levels; the presence of antiHBs in humans suggests protection from subsequent infections with the hepatitis B virus. This likely represents a species-specific hepadnavirus hepatitis. Since this report, similar clinicopathological findings have been seen in two Atlantic bottlenose dolphins (Bossart, unpubl. data).

Pinnipeds CASE 1 (Bossart and Schwartz, 1990)—Harbor Seal History

An adult, male harbor seal was examined because of acute anorexia and stereotypic swimming behavior. Moderate moist diffuse rates were auscultated in both lung fields. Mucous membranes were purplish-red and injected. Clinicopathological Findings 3

3

A marked leukocytosis (42,500 WBC/mm ; normal range for this seal: 7000 to 9000 WBC/mm ) 3 with absolute neutrophilia (39,525 neutrophils/mm ; normal range for this seal: 2660 to 6480


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neutrophils/mm ). The HCT was 73% (normal range 50 to 60%). The seal was hypernatremic (177 mEq/l; normal range: 147 to 156 mEq/l) and hyperchloremic (>130 mEq/l; normal range: 100 to 110 mEq/l). The BUN was 189 mg/dl (normal range: 44 to 60 mg/dl) and the LDH was >1500 mg/dl (normal range: 240 to 483 mg/dl). All other blood chemistry parameters were within normal ranges for this seal. Treatment

Amikacin sulfate was administered with intravenous 2.5% dextrose in half-strength lactated Ringer’s solution. Procaine penicillin G was administered IM. Nevertheless, the seal died. Post-Mortem Diagnosis

Acute necrotizing enteritis with positive fluorescent antibody staining of small intestinal tissue, with antisera to porcine transmissible gastroenteritis virus, feline infectious peritonitis virus, and canine enteric coronavirus, suggesting a coronavirus etiology. Discussion

Two other adult harbor seals housed with this seal died peracutely at the same time (Bossart and Schwartz, 1990). The histopathological findings in all three seals were similar. The absence of diarrhea in these cases may reflect the acute nature of this infection in seals. The source and transmission of this infection could not be determined. A feline source could not be ruled out, as feral cats were found at the facility prior to and during the disease outbreak.

Manatees CASE 1 (Walsh and Bossart, 1999; Walsh et al., 1999) History

Multiple orphaned manatee calves were hand-raised on different artificial milk formulas and developed a slow onset of anorexia, bloating, constipation, and/or diarrhea. Clinicopathological Findings 3

Calves typically had mild leukocytosis (generally not over 15,000 WBC/mm ) with absolute neutrophilia and lymphopenia. Fecal cytological examination generally indicated chronic–active or histiocytic inflammatory processes. Fecal cultures varied, indicating pure growths of Pseudomonas aeruginosa, Salmonella spp., Clostridium difficile (with or without toxin detection), Citrobacter freundii, or Escherichia coli. Abdominal radiographs sometimes indicated pneumatosis intestinalis (intramural intestinal accumulation of gas). Diagnosis

Enterocolitis, which was chronic–active, acute, histiocytic, hemorrhagic, and/or ulcerative. Treatment

Various regimens of oral antibiotic gut sterilization including gentamicin (2.5 mg/kg TID PO) and metronidazole (7 mg/kg BID PO) with bismuth salicylate, simethicone, and/or metoclopramide hydrochloride (see Chapter 43, Manatees) were utilized, as was elemental diet therapy (see Chapter 43, Manatees). Discussion

This condition in manatee calves can be life-threatening. The primary etiology may be related to the artificial nursing formula, immune system compromise (due to lack of a passive transfer of maternal immunoglobulins), and/or may represent nosocomial infections (see Chapter 43,


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Manatees). Notice the absence of a marked leukocytosis in spite of the severe life-threatening inflammatory disease. This is not uncommon in manatees.

Sea Otters CASE 1 (Rosonke et al., 1999) History

An adult, female Alaskan sea otter (Enhydra lutris lutris) rescued from the oil spill in Valdez, Alaska, developed an acute onset of caudal paresis after 8 years in captivity. The otter had no history of health problems prior to this episode. The otter was able to ambulate only by using its forelimbs. Clinicopathological Data

CBC, serum biochemical analysis (see exceptions), antibody titers against Toxoplasma gondii, serum antinuclear antibody, and urinalysis all were within normal ranges. The animal did have elevated CPK (2385 IU/l; reference range: 170 to 490 IU/l). Radiographic Results

Radiographs showed mild spondylosis of the lumbosacral vertebrae. Treatment

Dexamethasone, 0.2 mg/kg, BID PO, and trimethoprim–sulfamethoxazole, 20 mg/kg, BID PO beginning at day 10, were given. At approximately day 12, fasciculations of most skeletal muscles with postural ventral flexion of the neck at rest were noted with absence of a peripheral or deep pain response in both hind limbs. Bilateral flexor withdrawal reflexes were intact. The otter was unresponsive to humans, although appetite was normal. Compulsive grooming behavior was present. Further Clinicopathological Data

The CSF had a high nucleated count (46 cells/µl) and elevated RBC count (680 cells/µl). CSF protein = 36 mg/dl. CSF aerobic and anaerobic bacteria cultures were negative. Epaxial muscle biopsy indicated lymphoplasmacytic myositis with rhabdomyolysis and intracytoplasmic protozoan cysts consistent with Sarcocystis neurona by immunohistochemistry. CBC and serum analytes were normal. CSF and serum canine distemper titers were negative. Treatment

Pyrimethamine, 1 mg/kg, BID PO for 3 days then 1 mg/kg SID PO. Clinicopathological Data

A computed tomography (CT) scan of the brain was normal. Blood lead and mercury were within normal reference ranges for domestic animals. Clinical improvement was noticed in the next few days; however, within a week, paresis and increased ventriflexion of the neck were again detected. Magnetic resonance imaging (MRI) of the brain revealed high signal foci in the right dorsal pons and left thalamus, suggestive of demyelination. Serum and cerebral spinal fluid analysis for antibodies against S. neurona were positive. The clinical condition continued to deteriorate and the otter was euthanized 5 weeks after initial signs were noticed. Histopathological Diagnosis

The diagnosis was encephalomyelitis associated with a S. neurona-like organism. This was the first report of this condition in a sea otter (see Chapter 18, Parasitic Diseases).


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Acknowledgments The authors thank Sentiel Rommel for his drawings of venipuncture sites, Pam Yochem for her review of this chapter, Darey Shell for helping to compile the normal hematology and clinical chemistry tables, and Michelle Lander and Rebecca Duerr for editorial assistance.

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20 Cetacean Cytology Jay C. Sweeney and Michelle Lynn Reddy

Introduction Cytology, the microscopic study of cells, is a readily available inexpensive diagnostic tool and a valuable part of a preventive medical program. Similar to most nondomestic animals, marine mammals often mask early signs of poor health. However, disease processes often produce cytological abnormalities that, if examined, can indicate illness before the onset of obvious clinical signs (Cowell et al., 1999). Cytology may also be useful in monitoring the progress of a pathological process. Jergens et al. (1998) found a high correlation between the results of cytological and histological examination of samples collected by endoscopy of the stomach, small intestine, and colon of cats and dogs. Cytology in marine mammals is a developing field, with most samples examined to date having been taken from cetaceans rather than pinnipeds (Campbell, 1999). Husbandry programs at facilities that maintain captive marine mammals enhance their medical care by conditioning animals to allow physical examinations and collection of specimens without the need for physical restraint. Regular repetition maintains these behaviors, and allows for routine cytological monitoring. This provides baseline data for each animal, which is an important precursor for the effective evaluation of pathological cytology. Routine cytological examination facilitates the detection of many diseases in their early stages, thus allowing early implementation of therapeutic and control measures. The anatomy and physiology of cetaceans render cytological samples easily obtainable. In the fasted cetacean, the stomach almost always contains fluid, which acts as a repository for exfoliated cells that can be sampled and examined. The cetacean respiratory system is unique in that there is a rapid exchange of large volumes of air in a very short amount of time (5 l of air in 0.3 s). Cetaceans exchange about 80% of the volume of air in their lungs with a single breath, as compared with 20% or less in humans. Additionally, cetaceans lack nasal turbinate bones, which act as filters while exchanging air during inspiration and exhalation. As a result of these features, when a cetacean emits a “chuff ” characterized by a large volume of unfiltered, exhaled air traveling at high speeds directly from the deepest portions of the lung, a specimen of cell-rich exudate can be collected. Collecting samples from stranded cetaceans for cytological examination requires guidance from someone with experience in handling stranded animals. Precautions should be taken to protect both the animal and the rescuers from harm (Geraci and Lounsbury, 1993). In addition, care should be taken when handling and examining samples, as some fungi including Blastomyces, Histoplasma, and Coccidioides are potential airborne pathogens (see Chapter 17, Mycotic Diseases).

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If these fungi are suspected in the sample, care should be taken when handling the sample, including the use of a biological safety hood during slide preparation (Sweeney et al., 1976). The steps in cytological assessment are specimen collection, preparation, microscopic examination, and interpretation. Experience in examining cytological specimens will make interpretation easier. However, due to the number of factors that can affect the appearance of cells, interpretation may require consultation and confirmation with colleagues, reference laboratories, or a veterinary clinical pathologist. This chapter focuses on cytology of the respiratory tract, stomach, colon, rectum, and urinary tract of cetaceans, using the bottlenose dolphin (Tursiops truncatus) as an example. Relatively little is known about diagnostic cytology in other marine mammal species, but as more samples are examined baseline data will be established for them as well.

Sample Collection Modern husbandry programs incorporate conditioned animal behaviors that facilitate the collection of samples for health evaluation (Sweeney, 1984). Husbandry behaviors include, but are not limited to, ventral presentation for feces, urine, and milk collection; fluke or flipper presentation for blood collection; acceptance of a stomach tube for gastric sample collection; blowhole or nasal swab acceptance; and expulsion of sputum exudates (Sweeney, 1999). Because conditioned behaviors are performed voluntarily by the animal in its usual surroundings, samples may be collected frequently, and are less likely to reflect changes that may occur as an artifact of restraint. In stranded or otherwise unconditioned animals, sample collection is more difficult, but some samples, such as sputum, can still be collected rather easily, and perhaps opportunistically (Geraci and Sweeney, 1986). Specific methods for collecting samples from untrained animals are given in the chapters on individual groups of marine mammals (see Chapters 40 through 45). There are a few general guidelines for the collection of samples for cytology: 1. A small sample is usually sufficient; large volumes are not necessary. 2. To avoid contamination or distortion of the specimen due to desiccation or bacterial degradation, the sample should be evaluated as soon as possible after collection. 3. Specimens are best read within 6 hours of collection. If slides are to be stained with Papanicolauo (pap) stain, they should be fixed immediately (Head and Suter, 1975). 4. Avoid exposure of the sample to formalin fumes, alcohol, heparin, and/or excessive heat fixation, all of which can affect the quality of some stains.

Collection of Respiratory Tract Samples Sputum is collected from a conditioned cetacean by placing a sterile, nonabsorbable collection vial or petri dish over the blowhole and signaling the animal to produce several forceful exhalations or “chuffs” (Evers and Peddemors, 1986). Prior to specimen collection, the blowhole should be cleaned with an absorbent swab to remove water or other contaminants. If the animal is not conditioned for medical behaviors, a sample may be obtained by gently rolling the animal from side to side. This maneuver sometimes elicits a forced expiration (chuff), which expels exfoliated cellular material.

Collection of Gastric Samples When collecting a gastric sample, it is important to do so while the stomach is empty of food contents to eliminate the possibility of contamination of the specimen by food substances. Therefore, samples are best collected prior to the first feeding of the day. One common way to collect gastric fluid is by passing a 2-cm-diameter polyethylene tube (or smaller) directly into the stomach,


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applying mild suction upon entering the well of fluid in the first stomach compartment, then slowly removing the tube containing the fluid. It is also common to pass an endoscope into the stomach to collect gastric fluids for cytological examination and pH determination.

Collection of Fecal Samples Fecal samples can be collected from a conditioned dolphin as it positions belly up, parallel to the trainer’s platform. A soft, flexible 0.5-cm-diameter (or smaller) polyethylene tube is passed—up to 40 cm (15 in.) in a bottlenose dolphin—into the lower intestinal tract through the anal orifice. Fecal material is then gently aspirated into the tube. Note that the rectal mucosa is very delicate, and easily traumatized in cetaceans; therefore, care should be taken during specimen collection.

Collection of Urinary Tract Samples Urinary samples can be collected from a conditioned dolphin as it positions belly up near the trainer, or slides out in lateral recumbency and urinates on cue. The sample should be collected midstream. It can also be collected by catheter. For this, the animal is positioned belly up, parallel to the trainer’s platform. A soft, sterile flexible tube is placed through the urethral orifice (just caudal to the clitoris in females). The smallest tube that will permit collection should be used (usually a French gauge 8) and care should be taken to avoid trauma to the urethra or bladder.

Collection of Aspirates from Masses Fine-needle aspirates may be obtained from masses such as tumors or abscesses by techniques similar to those used in domestic dogs.

Slide Preparation With a basic medical-quality microscope operating according to specification, and the usual microscope supplies and various stains, a serviceable cytology laboratory can be developed. Some slides may be examined initially as wet mounts, whereas stains are necessary for retention and storage, and may be necessary for identification of certain conditions. Use the stain that is the easiest and quickest to use, while still providing sufficient cellular detail for identification. The basic materials and equipment needed for most cytological examinations are glass slides, coverslips, pipettes, wooden sticks or glass stirring rods, nonabsorbable vials, fecal flotation solution (e.g., sodium nitrate), litmus paper, and stains. Several basic stains are available, and details are given in standard texts, such as that of Boon and Drijver (1986). New methylene blue is quick and easy, whereas Wright–Giemsa-type stains are recommended for permanent slides, because they provide good cell differentiation. If Wright–Giemsa-type stains are used, the slide should be air-dried. For fungus detection, 10 to 20% potassium hydroxide, India ink, or lactophenol cotton blue are useful, while Gram’s stain is used for bacterial identification. If lactophenol cotton blue or Gram’s stain is used, the slide should be air-dried. Sudan stain is a standard for lipid staining, and pap stains work well, but only if the slides are fixed immediately (Head and Suter, 1975). Slide preparation methods will vary depending on the stain used; however, slides should be prepared as quickly as possible for best quality. When preparing a wet or dry mount, use a wooden stick, disposable pipette, or glass stirring-rod to mix one drop of sample and one drop of stain on a glass slide. If the sample is too viscous to produce a drop, the wooden stick can be used to transfer some of the sample to the slide. For a wet mount, cover the mixed


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FIGURE 1 For the dry mount technique, a small amount of sample is placed on a slide. A second clean slide is slid across the first at a 40° angle until it reached the edge of the drop (1), and then it is pushed back across the first slide. (From Sweeney, J.C., and Reddy, M., The Handbook of Cetacean Cytology, Dolphin Quest, San Diego, CA, 1999. With permission.)

FIGURE 2 For the squash technique, a small amount of sample is placed on a slide. A second clean slide is placed gently on top of the first (1), then pulled apart (2). (From Sweeney, J.C., and Reddy, M., The Handbook of Cetacean Cytology, Dolphin Quest, San Diego, CA, 1999. With permission.)

sample with a coverslip. For a dry mount, pull a second slide across the first at a 40° angle until it reaches the edge of the drop on the first slide. Then, gently, push it away again, across the first, spreading the sample across the first slide (Figure 1). Air-dry or fix, depending upon the stain used, following directions for the chosen stain. Evaluate slides microscopically using 10× and 40× objective lenses (for 100× and 400× magnification, respectively). Other optional treatments include centrifugation and the squash technique (Duncan and Prasse, 1986). Centrifugation may be necessary for samples with low cell concentrations. Although centrifugation will provide a greater number of cells for examination and identification, it does not allow for estimation of cell concentration and may damage or distort cells. In contrast, some specimens may require dilution with saline or water to facilitate visualization (e.g., some fecal specimens). The squash technique may be necessary for viscous samples (e.g., fibrin, mucus). For this technique, place a small amount of sample on the slide. Gently place another slide on top of the first, then pull them apart (Figure 2). Avoid putting too much pressure on the slides to prevent damaging cells. Stain. A useful method for examining feces is flotation. For this, place a small amount of fecal material in a nonabsorbable vial. Fill the vial with fecal flotation solution, place a coverslip on top of vial, and allow it to sit for 3 to 5 min. The specific gravity of saturated sodium nitrate may be less than seawater; therefore, alternative concentration techniques may be required to find some eggs and/or parasites. Fecal samples can also be prepared using sedimentation. For this, gently pour off the supernatant following examination of flotation material. Wash the sediment with fresh water and place a small portion on a microscope slide for examination.


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Examination of Specimens When examining the slide microscopically, it should be scanned under low power (100×) to detect giant cells, some parasites, and to assess the general composition of the slide. It may be necessary to use 400× for identifying specific cellular characteristics. Identification of some components, such as bacteria and fungi, may require using an oil-immersion objective.

Determination of Cellular Concentration within Slide Preparation Because considerable variation in sample concentration can occur between specimens, and at different areas of the slide preparation, it is most useful to record findings numerically as “mean values” or “ranges,” e.g., 5 to 10 epithelial cells per 400× field. Infrequent findings of significance (e.g., parasites, yeasts) can be noted as “rare” (one or two items observed throughout the entire slide), “occasional,” and/or in variable gradients such as 1+ to 4+.

Mucus Mucus in specimens is often highly cellular and may contain concentrations of leukocytes, which can accumulate as part of a reactive process. It is therefore helpful to record whether or not leukocytes are associated with mucus aggregates.

Amorphous Material Amorphous material is typical of most cytological preparations. When abundant, for example, in feces, it may be necessary to dilute the specimen to differentiate cellular material. Extraneous organisms, including saprophytic fungi, algae and diatoms, pollens, and parasitic larvae from dietary fish, are frequently encountered, especially in specimens from the gastrointestinal tract. Artifacts including dust on slides and precipitated or contaminated stain are also occasionally observed.

Interpretation Color Color of a sample prior to staining can be indicative of the following constituents: Clear to slightly yellow White Gray Green Brown Pink to red

A low concentration of cells Cells, mucus, fat droplets, chyle Cells, mucus, leukocytes Bile, bacteria, leukocytes Particulate debris, digested fish, bile, hemoglobin Erythrocytes, free hemoglobin

Epithelial Cells Epithelial cells are often present in properly collected specimens. The type of epithelium is reflective of the anatomical site from which it is collected—e.g., squamous (gastric, respiratory, lower urinary tract, mammary, i.e., large ducts), columnar and/or secretory (gastric, respiratory, colorectal, mammary), cuboidal (renal tubular, bladder, mammary, i.e., ductal and secretory cells), and transitional (urinary bladder). This chapter deals primarily with exfoliative cytology, where specimens to be examined originate from an organ system with lumenal


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surfaces lined by epithelial cells. In such samples, the number of epithelial cells generally serves as a good index of specimen concentration.

Leukocytes Leukocytes (white blood cells, wbc) are indicative of inflammation, which is often associated with infection and/or necrosis. The presence of leukocytes does not necessarily signify disease. In the experience of the authors, a usable measure of the relative number of leukocytes present is the ratio of leukocytes to epithelial cells. When the ratio is greater than or equal to 1 : 1, then ongoing inflammation and/or necrosis is a reasonable assumption.

Erythrocytes The presence of erythrocytes (red blood cells, RBC) is indicative of hemorrhage or diapedesis. However, careful consideration must be given to method(s) used to acquire specimens. For example, when colorectal specimens are obtained via rectal catheterization, or via rectal swabs, mucosal trauma may result in incidental bleeding into the specimen.

Respiratory Tract Normal Findings The cells found in a cetacean sputum sample may originate from lung, trachea, bronchi, nasal sacs, pterygoid sinus, or the larynx. Normal findings and reference values for respiratory samples are shown in Table 1. Start with an epithelial cell count when examining a respiratory sample. Be aware that the appearance of these cells can vary depending on their orientation on the slide. Squamous epithelial cells (Color Figures 1E and 1H)* are commonly found in respiratory samples and are generally used as an index to assess sample concentration and the significance of other findings. For example, the presence of a few leukocytes (Color Figure 1P) is considered normal as long as there are fewer of them than epithelial cells. TABLE 1 Normal Cytological Findings and Their Reference Ranges in Samples from Bottlenose Dolphins (Tursiops truncatus) (numbers are means per field) Feature

Magnification, ×

Respiratory Tract

Stomach

Epithelial cells

400

>5

>5

Macrophages Leukocytes Degenerated leukocytes Erythrocytes Eggs Protozoa Fungi Casts

400 400 400

0 0–5 0

400 100 100 400 400

0 0 0–1+ 0 0

*Color Figures follow p. 462.

Feces

Urine 2–10

0–2 0–5 0

0 to too many to count 0 0–5 0–10

0 0 0 0 0

0–3 0 0 0 0

0–2 0 0 0 0–2

0 0–2 0


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Some common contaminants in respiratory samples include stain precipitate from old stain, salt crystals, powder from latex gloves, fibers from collection swabs, diatoms, algae, and pollen, which can be mistaken for helminth eggs or protozoan cysts. It is a good idea to become familiar with the diatoms, algae, and/or pollen common to the area where samples are collected to help in the identification of these contaminants. The significance of some items such as bacteria should be considered in context with cellular evidence of inflammation. In the absence of inflammation, bacteria such as Simonsiella (Color Figure 1D) are considered to be normal flora in a sputum sample. Similarly, the respiratory tract of a healthy cetacean may contain Candida (Color Figure 1L). If it does not invade healthy tissue, and as long as the animal’s immune system is not compromised, its presence is most likely not a clinical problem (see Chapter 17, Mycotic Diseases) (Dunn et al., 1982; Haebler and Moeller, 1993). Similarly, light infections of parasites such as holotrich ciliates are considered common findings in cetaceans. For example, Kyaroikeus cetarius (Sniezek et al., 1995; Color Figure 1I) is commonly found in the blowhole of bottlenose dolphins, and is found in more than 50% of free-ranging animals (Woodard et al., 1969). It may be the only facultative endoparasite of marine mammals (Geraci and St. Aubin, 1987). Similarly, if found in the absence of erythrocytes or leukocytes, Nasitrema eggs may not be associated with pathology (Sweeney, 1986).

Significant Findings A leukocytes-to-epithelial cell ratio greater than 1 in sputum or the presence of macrophages (Color Figure 1A) suggests inflammation. Band cells, which are immature neutrophils, usually appear in response to acute infectious or inflammatory conditions. The nucleus looks like a curved band (Color Figure 1G). Another indicator of inflammation is fibrin (Color Figure 1B). Erythrocytes (Color Figure 1F) indicate gross or microscopic hemorrhage. The eggs of the trematode Nasitrema sp. in the presence of erythrocytes or leukocytes can indicate damaged capillaries and/or inflammation. Nasitrema or other upper respiratory infections are often seen in conjunction with heavy infections of ciliates such as K. cetarius (Sniezek et al., 1995; Color Figure 1I). Active infection with the lungworm Halocercus may result in larval forms in sputum samples (Woodard et al., 1969; Sweeney and Ridgway, 1975; Haebler and Moeller, 1993). There are many bacteria (Color Figure 1K) found in the lungs of cetaceans with bronchopneumonia (see Chapter 16, Bacterial Diseases) (Sweeney and Ridgway, 1975; Sweeney, 1978; Howard et al., 1983). Aspergillus spp. (see Color Figure 1O) are observed in sputum samples associated with aspergillosis (see Chapter 17, Mycotic Diseases) (Sweeney et al., 1976; Migaki and Jones, 1983; Carroll et al., 1986; Joseph et al., 1986). Candida spp., a small, budding yeast (see Color Figure 1L), is often present, but when invasive it often forms pseudohyphae. To assess pathogenicity, it is important to note the progression in abundance through successive specimens (Medway, 1980; Dunn et al., 1982; Reidarson et al., 1996). A significant fungal pathogen occasionally observed causing necrotizing cutaneous and muscular lesions in cetaceans is Apophysomyces elegans (Cunninghamella sp.), an opportunistic Zygomycete fungus (see Chapter 17, Mycotic Diseases; Robeck et al., in press; Color Figure 1M).


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Stomach Normal Findings Before preparing slides of gastric samples for cytological examination, note the color, consistency, odor, and pH. The pH of normal gastric fluid from a fasted dolphin can vary from as low as 1.5 to as high as 3.0. Care must be taken to avoid contamination with seawater prior to measuring pH. Low pH will lyse erythrocytes, so these may be missed when examining a gastric sample for cytology. To overcome this, one may deliver a neutral solution (e.g., saline) prior to collection of the sample for cytology, then immediately aspirate the solution and examine the sediment, or simply add a pH 7 buffer to a sample as soon as it is collected. It is important to measure the pH of a sample prior to examining it, to be able to interpret cytological findings. Clinically significant changes in gastric pH are discussed in Chapter 40, Cetaceans. Reference values for cytological samples collected from the stomach are given in Table 1. The cells found in a gastric fluid sample may originate from the oral mucosa, larynx, esophagus, stomach, or the respiratory tract. At low pH conditions, many epithelial cells are crenated and leukocytes exhibit varying degrees of cell membrane and cytoplasm loss, leaving only the nucleus. The degree of leukocyte staining thus varies. The acid conditions typically lyse erythrocytes so they cannot be visualized cytologically. In cases of overt gastric bleeding, specimens appear grossly brown to red-brown, which also produces a strong positive occult blood test. Note that gastric and fecal specimens typically exhibit or test positive for occult blood as a result of fish blood in the diet. Squamous epithelial cells found in a gastric sample are likely to have originated in the upper gastrointestinal tract. The significance of bacteria (Color Figure 1K) should be considered in context with cellular evidence of inflammation. Although yeasts are an occasional normal finding, they may be significant if abundant. Some findings in a sample are a result of a fish diet. For example, food fish are typically infested with adult and larval nematodes, which may therefore appear in a stomach sample, as may oil droplets. However, if fish particles are present in a gastric sample following an overnight fast, maldigestion should be considered. Amorphous debris is normal and will be minimal if the gastric sample is collected prior to the animal’s first meal of the day.

Significant Findings The presence of many leukocytes (Color Figure 1P) and basal cells (Color Figure 1N) from the gastric submucosa may suggest gastric erosion or ulceration, especially if leukocytes are not also found in the sputum. Long-lived macrophages (Color Figure 1A) are suggestive of chronic inflammatory lesions. Band neutrophils (Color Figure 1G) suggest response to an infectious or inflammatory condition. No erythrocytes (Color Figure 1F) should be visible in a gastric sample because of rapid lysis when exposed to the acidic pH of gastric fluid. Their presence indicates a significant increase in gastric pH, and suggests gastric ulcers with bleeding, especially in the presence of leukocytes (Color Figure 1P). Nasitrema eggs may be seen in the gastric sample if they are swallowed from the respiratory sputum (Dailey and Stroud, 1978; Howard et al., 1983). Eggs from the trematode Braunina cordifromis are found in the fundic chamber of the stomach. Damage caused to gastric mucosa is minimal (Schryver et al., 1967; Howard et al., 1983; Haebler and Moeller, 1993).


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Colon/Rectum Normal Findings The cells in a colon or rectal sample may originate from the respiratory tract, stomach, duodenum, intestine, or anus. Normal findings and reference values are shown in Table 1. As in respiratory and gastric samples, epithelial cells (columnar and squamous; Color Figure 1H) are used as an index to determine the concentration of the colon/rectal sample, and the number of white blood cells (Color Figure 1P) should be considered in context with other cellular processes. Mucus in specimens is often highly cellular. If present, digested fish particles may be difficult to identify in a sample from a healthy animal; however, lipid droplets as a result of a fish diet are an expected component of a normal fecal sample. Bacteria (Color Figure 1K) are also common in a fecal sample. Some common contaminants in colon and rectal samples include stain precipitate from old stain, powder from latex gloves, and pollen. Pollen shapes vary depending on the plant from which they originate, so becoming familiar with pollen common to the area may facilitate identification.

Significant Findings Too numerous to count (TNTC) degenerated leukocytes indicate inflammation in a colon/ rectal sample. A fecal leukocyte count may be used to aid diagnosis of enteritis (Sweeney and Ridgway, 1975). In the presence of leukocytes, erythrocytes (Color Figure 1F) are significant. Erythrocytes are not always observed in cases of intestinal bleeding, as they may be lysed prior to excretion, and occasionally are not passed to the colon if intestinal stasis is severe. Undigested fish particles in a fecal sample may indicate improper digestion. Budding yeasts or fungal hyphae in a fecal sample (Color Figure 1J) could be a result of intestinal candidiasis, or other fungal infection. Campula rochebruni is found in the hepatic and pancreatic ducts, and has been implicated in cases of colicystitis and pancreatitis (see Sweeney and Ridgway, 1975). Other parasitic eggs and larvae can also occur in fecal specimens (see Chapter 18, Parasitic Diseases).

Urinary Tract Analysis of urine should be done as quickly as possible after collection, because chemical and cytological changes occur rapidly, especially if the sample is kept at room temperature. Bacterial growth can increase the pH, and in alkaline urine, casts tend to dissolve, and may disappear in time. Refrigerated samples should be warmed to room temperature before examination. Before preparing the slide, the sample should be gently agitated to resuspend the sediment. Use low light to examine unstained sediments, and to facilitate seeing elements such as casts.

Normal Findings A wide variety of epithelial cells can be found in the urine and can come from the bladder, urethra, renal pelvis, or ureters. Squamous epithelial cells (Color Figure 1H) are the largest cells seen in a normal urine sample. Thin and flat, they may be present as single cells or in small clusters. If they are rolled, they may look like casts. Commonly found in lower numbers in voided samples, they are usually a result of genital tract contamination. Transitional epithelial cells (Color Figure 1C) originate from the renal pelvis, ureters, urinary bladder, and/or urethra. They differ in size and shape depending on their locations in the mucosa. Generally, they are


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smaller and have smoother edges than squamous epithelial cells and are larger than leukocytes (Color Figure 1P). There are usually only a few single cells or small clusters in voided samples, but there may be more, perhaps sheets, in samples collected by catheterization. Goblet cells are columnar epithelial cells that produce mucus and may be found in the prostate, the vaginal vault, or within the urethra. Leukocytes and erythrocytes (Color Figures 1F and 1P) are found in normal urine as a result of diapedesis in the urinary tract, and a few red blood cells (up to five at 400×) may be found in a normal sample collected by catheter. Spermatozoa are a common finding in samples collected from males. Normal urine is sterile in the bladder, but may become contaminated with small numbers of bacteria (Color Figure 1K) as the urine is voided. Mucus cells may protect against bacterial infection and are a common finding in a urine sample. Casts are cylindroid proteinaceous structures formed in the renal tubule lumen. In a normal sample, there are few to none. Crystals are rarely seen, and their clinical significance is unclear.

Significant Findings Renal problems are generally rare in cetaceans (Sweeney, 1986). Although a variety of epithelial cells may be found in the urine, the number of such cells is increased in animals with cystitis or infections in the urogenital tract. Numerous leukocytes in a urine sample indicate a pathological process somewhere along the urinary (or urogenital in voided specimens) tracts. A combination of erythrocytes and leucocytes (Color Figures 1F and 1P) may suggest infection or trauma (e.g., cystitis, nephritis). Erythrocytes without leukocytes may suggest hemorrhage due to urinary calculi, trauma, or neoplasia somewhere along the urinary tract (or urogenital tract in voided specimens). Erythrocytes are smaller than leukocytes and may appear colorless to yellow or orange and do not stain with new methylene blue. In conjunction with large numbers of leukocytes, large numbers of bacteria (Color Figure 1K) may be seen in animals with cystitis or bacterial infections elsewhere in the genital and urinary tracts. Increased numbers of casts in urine sediment are usually a result of damage to renal tubular cells. Progressive urinary tract infections result in fixed specific gravity and casts of the waxy or fine type (Small, 1975). These may be confused with rolled epithelial cells. Yeasts (Color Figure 1J) may be present in a urine sample as a contaminant, and this should be suspected if the sample is voided and/or old. If the sample is fresh, fungal infection of the kidneys and/or bladder should be suspected.

Acknowledgments The authors thank marine mammal trainers everywhere, who have been involved in conditioning the husbandry behaviors that allow for the collection of cytological samples, and those dedicated to rescuing and rehabilitating wild marine mammals. Thomas Lipscomb, John Bjorneby, and Sam Ridgway are acknowledged for their expertise in cytology, and Judy Lawrence and Howard Rhinehart are also thanked for reviewing this chapter.

References Boon, M.E., and Drijver, J.S., 1986, Routine Cytological Staining Techniques: Theoretical Background and Practice, Macmillan, Houndmills, England, 238 pp. Campbell, T.W., 1999, Diagnostic cytology in marine mammal medicine, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 464–469. Carroll, J.M., Jasmin, A.M., and Bascom, J.N., 1986, Pulmonary aspergillosis of the bottle-nosed dolphin (Tursiops truncatus), Am. J. Vet. Clin. Pathol., 2: 139–140.


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Cowell, R.L., Tyler, R.D., and Meinkoth, J.H., 1999, Diagnostic Cytology and Hematology of the Dog and Cat, C.V. Mosby, St. Louis, MO, 338 pp. Dailey, M.D., and Stroud, R., 1978, Parasites and associated pathology observed in cetaceans stranded along the Oregon coast, J. Wildl. Dis., 14: 503–511. Duncan, J.R., and Prasse, K.W., 1986, Veterinary Laboratory Medicine: Clinical Pathology, 2nd ed., Iowa State University Press, Ames, 285 pp. Dunn, J.L., Buck, J.D., and Spotte, S., 1982, Candidiasis in captive cetaceans, J. Am. Vet. Med. Assoc., 181: 1316–1321. Evers, P., and Peddemors, V., 1986, A scanning electron microscope study of a ciliate obtained from dolphin blowholes, Electr. Microsc. Soc. S. Afr. Proc., 16: 33–34. Geraci, J.R., and Lounsbury, V.J., 1993, Marine Mammals Ashore: A Field Guide for Strandings, Texas A&M University Sea Grant College Program, College Station, 305 pp. Geraci, J.R., and St. Aubin, D.J., 1987, Effects of parasites on marine mammals, Int. J. Parasitol., 17: 407–414. Geraci, J.R., and Sweeney, J.C., 1986, Clinical techniques, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 771–777. Haebler, R., and Moeller, R.B., Jr., 1993, Pathobiology of selected marine mammal diseases, in Pathobiology of Marine and Estuarine Organisms, Advances in Fisheries Science, Couch, J.A., and Fournier, J.W. (Eds.), CRC Press, Boca Raton, FL, 217–244. Head, J.R., and Suter, P.F., 1975, Approach to the patient with respiratory disease, in Textbook of Veterinary Internal Medicine: Diseases of the Dog and Cat, Ettinger, S.J. (Ed.), W.B. Saunders, Philadelphia, 544–564. Howard, E.B., Britt, J.O., and Matsumoto, G.D., 1983, Parasitic diseases, in Pathobiology of Marine Mammal Diseases, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 11, 96–162. Jergens, A.E., Andreasen, C.B., Hagemoser, W.A., Ridgway, J., and Campbell, K.L., 1998, Cytologic examination of exfoliative specimens obtained during endoscopy for diagnosis of gastrointestinal tract disease in dogs and cats, J. Am. Vet. Med. Assoc., 213: 1755–1759. Joseph, B.E., Cornell, L.H., Simpson, J.G., Migaki, G., and Griner, L., 1986, Pulmonary aspergillosis in three species of dolphin, Zoo Biol., 5: 301–308. Medway, W., 1980, Some bacterial and mycotic diseases of marine mammals, J. Am. Vet. Med. Assoc., 177: 831–834. Migaki, G., and Jones, S.R., 1983, Mycotic diseases in marine mammals, in Pathobiology of Marine Mammal Diseases, Vol. 1, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 1–127. Migaki, G., Blumer, P.W., and Augsburg, K., 1975, Case for diagnosis: Phycomycosis in a dolphin, Mil. Med., 140: 544–549. Reidarson, T.H., McBain, J., and Harrell, J.H.,1996, The use of bronchoscopy and fungal serology to diagnose Aspergillus fumigatus lung infection in a bottlenose dolphin (Tursiops truncatus), Abstr., Proceedings of the 27th International Association for Aquatic Animal Medicine, Chattanooga, TN, 34. Robeck, T., Dalton, L., and Rinaldi, M., in press, Zygomycosis infections in a bottlenose dolphin (Tursiops truncatus), killer whale (Orcinus orca), and two Pacific white-sided dolphins (Lagenorhynchus obliquidens) caused by Saksenaea vasiformis and Apophysomyces elegans. Schryver, H.F., Medway, W., and Williams, J.F., 1967, The stomach fluke, Braunina cordiformis, in the Atlantic bottlenose dolphin, J. Am. Vet. Med. Assoc., 151: 884–885. Small, E., 1975, The Mycoses, in Textbook of Veterinary Internal Medicine: Diseases of the Dog and Cat, Ettinger, S.J. (Ed.), W.B. Saunders, Philadelphia, 181–202. Sniezek, J.H., Coats, D.W., and Small, E.B., 1995, Kyaroikeus cetarius n. g., n. sp.: A parasitic ciliate from the respiratory tract of odontocete cetacea, J. Eukaryotic Microbiol., 42: 260–268. Sweeney, J., 1978, Infectious diseases of body systems, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 589–592. Sweeney, J.C., 1984, Medical procedures—The easy way, Soundings, 9: 2. Sweeney, J., 1986, Clinical consideration of parasitic and noninfectious diseases, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 785–789.


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Sweeney, J.C., 1999, How good is that blow sample? Soundings, 24: 9. Sweeney, J.C., and Reddy, M., 1999, The Handbook of Cetacean Cytology, Dolphin Quest, San Diego, CA, 1999. Sweeney, J.C., and Ridgway, S.H., 1975, Common diseases of small cetaceans, J. Am. Vet. Med. Assoc., 167: 533–540. Sweeney, J.C., Migaki, G., Vainik, P.M., and Conklin, R.H., 1976, Systemic mycoses in marine mammals, J. Am. Vet. Med. Assoc., 169: 946–948. Woodard, J.C., Zam, S.G., Caldwell, D.K., and Caldwell, M.C., 1969, Some parasitic diseases of dolphins, Vet. Pathol., 6: 257.


21

Gross Necropsy and Specimen Collection Protocols Teri K. Rowles, Frances M. Van Dolah, and Aleta A. Hohn

Introduction Data and specimens can be collected from both live and dead marine mammals, and are vital for the determination of individual animal health and for studies of the biology and health of populations of these animals, both in captivity and in the wild. The specific goals of scientists, managers, and veterinarians for data and specimen collection may vary. However, all aspects of collection and interpretation will benefit from standardized collection protocols and from data sharing for assessments. The types of assessments extracted from the data and specimens collected can include cause of death or illness, success of management practices, life-history shifts or definitions, the presence and effects of noise, chemical pollutants, new pathogens, and/or ecological changes on overall population health, recovery, or decline. Although it may appear that these assessments are relevant only to wild populations, they are also important in both the management of populations in public display facilities and the conservation measures undertaken with critically endangered species through captive management. Three factors are important for the interpretation of the data obtained from assessments and specimen analyses: the development and use of standardized collection protocols; the use of quality assurance methods in specimen collection and analyses; and the sharing of data and protocols among regional, national, and international groups. This chapter is intended for veterinarians, veterinary students, researchers, and biologists who are working with either captive marine mammals or marine mammals in the wild. Although most of the chapter focuses on the collection of data and specimens through necropsy examinations, the collection of data and specimens from live animals is also important in public display or wild population management assessments. In the United States, much of the health assessment for wild populations of marine mammals is guided by the Marine Mammal Health and Stranding Response Program established by the Marine Mammal Protection Act (see Chapter 5, Unusual Mortality Events; Chapter 33, Legislation), and therefore scientists and managers have made strong efforts to coordinate health assessment protocols and to share information. In other countries there are similar efforts to nationally and regionally standardize the types of information collected and the analytical methods used, especially through international

0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

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organizations such as the International Whaling Commission (IWC), the International Council for Exploration of the Seas (ICES), and the Arctic Monitoring and Assessment Program (AMAP).

Necropsy Examinations and Specimen Collection Information on morbidity and mortality is scarce for many wildlife populations, including marine mammals. Although efforts over the last decade have increased the knowledge of certain pathogens (i.e., morbillivirus) in marine mammals (Duignan et al., 1996), many data gaps still remain. To better understand the overall health of marine mammal populations, examinations of carcasses from a variety of sources, such as strandings, subsistence hunts, or incidental fishery by-catch, are needed. These interrelationships are best examined through multidisciplinary studies, which benefit from standardized protocols. The goals of standardized necropsy examinations are to accomplish the following: • • • • • •

Determine cause of death; Collect basic biological data; Determine direct human impacts (e.g., fishery by-catch; ship strike); Collect data for management assessment; Establish baselines of health, disease, and biology; Understand the levels of exposure and the effects of biotoxins, chemical pollutants, pathogens, noise, and other environmental factors on health.

Among other factors (see Chapter 5, Unusual Mortality Events; Chapter 4, Stranding Networks), in response to the 1987–1988 bottlenose dolphin (Tursiops truncatus) die-off along the Atlantic Coast of the United States, the Marine Mammal Health and Stranding Response Program was established. One of the major goals of this program is to coordinate a more effective response to marine mammal unusual mortality events (MMUMEs). To that end, the National Marine Fisheries Service (NMFS) and the U.S. Fish and Wildlife Service (FWS) have developed contingency plans to guide the investigations of UMEs, with consultation from a group of advisors (the Working Group on Marine Mammal Unusual Mortality Events; see Chapter 5, Unusual Mortality Events) (Wilkinson, 1996; Geraci and Lounsbury, 1997). If one is investigating mortalities of stranded animals that appear unusual in number or condition, one must immediately notify the regional or species stranding coordinator, who then should take immediate steps necessary to consult with the working group described above. Even before an event has been declared “unusual” (a formal designation), additional or new protocols for each species may be necessary. Once an event has been formally designated as an unusual event, protocols for the investigation will be provided by the on-site coordinator and the Working Group on Marine Mammal Unusual Mortality Events. The stranding network personnel will be critical in the response to the event and in the interpretation of the data obtained from in-depth analyses. Comparisons of data collected from unusual events with data obtained during routine necropsies will enhance the interpretation and assessment of each event investigation. Carcasses may not always be in an optimal state for all protocols, depending on the stage of decomposition, but some information can be gained from carcasses in all states of decomposition. Throughout this chapter, references will be made to the standardized carcass decomposition categories for the U.S. National Stranding Network (Geraci and Lounsbury, 1993) (Table 1). As with other species protected by law, determination of cause of death may lead to litigation or prosecution. Therefore, the collector should follow standardized protocols, document field collections, use and document photography, and in certain instances maintain chain of custody for all samples and data collected during an investigation (Figure 1).

Live

Freshly dead “edible”

Moderate decomposition

Advanced decomposition

Severe decomposition

2

3

4

5

Definition

1

Code

Gross Appearance

No bloating; minimal drying and wrinkling of epidermis in cetaceans and manatees or dermis and epidermis in pinnipeds and otters, and of eyes and mucous membranes; muscles firm; blubber firm and white or yellow; internal organs intact; liver still with physical integrity Slight bloating with tongue and penis protruding; some skin sloughing and cracking; eyes sunken; blubber may be bloodtinged; muscles soft; all internal organs including liver still have gross integrity but are soft and friable Bloated; missing patches of epidermis and hair; internal organs show lack of integrity and are extremely friable; blubber with gas pockets and pooled oil Mummified; skeletal

TABLE 1 Classification of Carcass Condition

Cause of death only rarely determined

Autolysis often masks cause of death; bloating and autolysis may alter morphometrics

Morphometrics, gross pathology, parasitology, genetics, life history

Limited morphometrics, age, skeletal pathology, genetics

Autolysis often masks histological assessment; decomposition may alter enzymatic, biochemical, and chemical analyses, including lipid quality and quantity

Bacterial overgrowth may be observed on cultures or histology; some autolysis noted on histology

Interpretation

Morphometrics, gross pathology, parasitology, genetics, life history, some histology

Morphometrics, blood, biopsies, urine, infectious diseases, diagnostic imaging All types of specimens should be collected

Specimen Collection


Gross Necropsy and Specimen Collection Protocols 451


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Figure 1 Chain of Custody Form. This form is used to track specimen sample transfers for marine forensic studies. It ensures that one is always aware of where a sample is at any particular time, should it be needed for additional testing or legal examination. Note that the form is signed and dated both on receipt and on release of the specimen. (Form courtesy of National Marine Fisheries Service.)


Gross Necropsy and Specimen Collection Protocols

453

Carcass Condition Code Carcasses are rated as to the state of decomposition on a scale of 2 to 5 (Code 1 = alive) (see Table 1 for code descriptions). The condition code will limit the measurements taken, but some information can be gained from animals at all stages of decomposition. The carcass condition codes that are appropriate for each type of sample are listed in Tables 1 through 4.

Morphometrics A standard set of morphometric and descriptive data should be collected on all marine mammal carcasses, and on any marine mammal that is captured for assessment, research, or tagging. These data provide information important for understanding the basic biology of specific marine mammals, basic stock structure, demographic trends, nutritional status, population trends, and epidemiological investigations of diseases, mortality events, or human interactions. Morphometrics can be used to assist with species identification, age-class estimation, and body condition. Determination of body condition, age class, and reproductive class is required for the interpretation of pollutant burdens and effects in marine mammals, and for epidemiological interpretation of diseases and mortality events. Whenever possible, skulls of stranded animals should be collected as voucher specimens or archival specimens, particularly in mass strandings or die-offs. Morphometric Data Protocol

For each taxon there are specific measurements that should be taken. Measurements should be standardized, and examinations and measurements should be augmented by photography. Each photograph should include the animal identification, date, and some means of assessing measurements (e.g., a xeroxable scale on the identification label). Photographs, like straightline measurements, should be taken so that size, shape, and position can be obtained. Girth measurements may or may not be possible or useful given the size and posture of the animal, the state of decomposition, and location of the carcass. However, some estimation of girth may be possible (e.g., measuring the distance from dorsal midline to ventral midline and multiplying by two). Blubber measurements should be taken from specific locations in each taxon, as they are important for assessment of nutritional status and overall body condition. Multiple blubber depth measurements should be taken, since blubber thickness varies with body region. Blubber assessments may be rapidly affected by decomposition, exposure, and autolysis, including such aspects as depth, amount of lipid (due to leaching), and types of lipid classes (Krahn et al., in press). Body organs or body compartments should be weighed whenever possible. However, autolysis and/or dehydration (freezing artifacts) may alter weights and measurements and must be so noted. Organ weights from carcasses in moderate to advanced decomposition states or from carcasses frozen for long periods of time may not be useful and may weaken organ weight databases.

Genetics Knowledge of the species, as well as the specific population from which an animal came, is critical for interpreting data collected from live or dead animals. Many different tissues have been used for genetic analysis; however, skin and liver are the most commonly collected tissues. White blood cells, muscle, gonads, teeth, and bone have also been collected from carcasses, and white blood cells or skin biopsies are typically collected from live animals (see Chapter 14, Genetics).


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Genetic Sample Protocol

Genetic analyses require only a small sample; the recommended sample size for collection is 3 3 0.5 cm (∼0.2 in. ) soft tissue cut into small strips for fixation; 1 ml of whole blood, whole teeth, or a piece of bone have also been collected for genetics. The best method of preservation depends on the tissue collected. Soft tissue, such as skin, is best preserved in 5 to 20% DMSO in saturated salt solution at 1 volume of tissue to 10 to 20 volumes of preservative. The solution containing the tissue should then be frozen for long-term storage. DNA can be extracted from frozen soft tissue without preservative, but it is more difficult, particularly if nuclear DNA (e.g., microsatellites) is to be analyzed. Alternative methods include fixation in 80% ethanol or drying. Blood samples are best frozen at –80°C or colder.

Stomach Contents Evaluation of stomach contents is important both for diagnostic evaluation and for biological assessment of prey selection. Stomach content analyses are time-consuming efforts and should be performed by experienced personnel; however, collection and storage of contents are easy to perform in the field. Stomach contents may include otoliths, macerated prey flesh, skeletal remains, parasites, foreign bodies, and vegetation. Fish otoliths are one of the most commonly used structures for prey identification. The shape and characteristics of an otolith are species specific, and the size of the otolith is proportional to fish size, allowing for evaluation of size class of prey as well as caloric intake of the marine mammal. Stomach Contents Protocol

In small animals, the stomach may simply be tied off at both ends and frozen intact for later examination, although freezing may limit pathological and parasitological examinations. Ideally, the stomach should be opened when fresh, the mucosa gently flushed with saline, and the contents (including the washings) frozen or fixed for later evaluations. The type of preservation of the contents will depend on the expected diet of the various taxa of marine mammals. Buffered neutral formalin fixation may dissolve the otoliths of some prey fishes; therefore, formalin fixation should not be used for preserving stomach contents from fisheating marine mammals (Geraci and Lounsbury, 1993). These contents instead should be frozen or fixed in alcohol. Stomach content samples from plant-eating marine mammals (e.g., manatees) should not be frozen, since the freezing of seagrass and algae causes fragmentation of the cells, making identification very difficult (Eros et al., 2000). Instead, stomach contents from herbivorous animals should be preserved in 5 to 10% neutral buffered formalin or 80% ethanol at a ratio of 1:1 or 2:1 (Eros et al., 2000; Rommel, pers. comm.). Subsamples for toxicology or biotoxins are collected from the stomach contents of fresh carcasses when they are first opened, and the subsamples frozen for later evaluation. If the stomach is opened fresh, parasites can be collected (see below) and the mucosa examined for pathology. Freezing and thawing may limit identification and interpretation of gastrointestinal pathology and parasites. Foreign bodies should be documented and photographed, and ingested marine debris or fishing gear saved whenever possible.

Age Estimation of age for specific animals, or within specific stranding events, is important from an epidemiological perspective, as well as important in understanding the basic biological characteristics of a particular species or stock. Currently, age is estimated primarily from counts of growth layers deposited in several persistent tissues, primarily teeth and, less often, bone.



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Growth layers in these persistent structures are similar in concept to growth rings in trees. Saving teeth or other tissue for aging from known-age animals (from the wild or captive situations) is also important, because these tissues are used to validate the interpretation of growth layers for specific taxa. At times, relative measures of age, such as tooth wear, pelage or skin color, or fusion of cranial sutures, which allow individuals to be placed in age groups, are helpful. Age class or maturation status may be estimated using body size (length) (Stevick, 1999), fusion of epiphyses, pelage color, or reproductive parameters. The use of body size as a rough estimate of age, however, requires that a growth curve has been generated for that species from running models that fit size-at-age data for a large number of specimens whose age was known or estimated from growth layers. Growth layers (or growth layer groups; Perrin and Myrick, 1980) in teeth have been used to estimate age for odontocetes and pinnipeds (Hohn et al., 1989; Oosthuizen, 1997), since they were first associated with age by Scheffer (1950). For small cetaceans, growth layers are counted primarily in dentine, although for a few species (e.g., the fransciscana (Pontoporia blainvillei) and beaked whales) cement is better. For pinnipeds, growth layers are counted in both dentine (the yellowish, calcified tissue that makes up the bulk of all teeth, harder than bone, softer than enamel) and cement (thin, bonelike material covering roots of teeth, softer than dentine). Canines are best for dentinal counts, but in very old animals the pulp cavity may be occluded, and cement must then be used. Cement is best counted in post-canines (Klevezal, 1996). Incisors can be safely extracted from live animals, but these smaller teeth have small layers, and age tends to be underestimated by significant amounts in old animals (Bernt et al., 1996). For dugongs, the tusk (incisor) or canine can be used (Eros et al., 2000). For a number of species, notably manatees and baleen whales, teeth cannot be used for age estimation. Manatees have an indeterminate number of molars that are constantly lost and replaced throughout life, and no tusks. Baleen whales have no teeth. Fortunately, annual growth layers do occur in the tympano-periotic (auditory) bones of manatees (Marmontel et al., 1996) and baleen whales (Klevezal, 1996). For each species, the location on the bone with the maximum number of layers must be found; in other regions, resorption of early-deposited layers results in an underestimate of age. In all bones, growth layers occur in periosteal bone and generally the maximal number of layers occurs where the periosteal bone is thickest. In the balaenopterid whales, earplugs also have been used for age estimation (Lockyer, 1984; Kato, 1984). These structures are actually a horny epithelium formed in layers on the external surface of the tympanic membrane of the external auditory meatus. In addition to numbers of growth layers, a change in the morphology of the growth layers from irregular layers (immature) to regular layers (mature) has been seen in some species, and is thought to indicate the transition to maturation (Thomson et al., 1999). Chemical signals, specifically amino acid racemization, have been used for dolphins and small and large species of whales (Bada et al., 1980), including, most recently, fin (Balaenoptera physalus) and bowhead whales (Balaena mysticetus) (George et al., 1999). Age is estimated as a function of the proportion of d- and l-isomers of aspartic acid in the lens of the eye. Accurately and precisely counting the annual layers depends greatly on the tissue and techniques used. For example, Hohn and Fernandez (1999) found that stained sections allow more accurate estimates of age in bottlenose dolphins, and Stewart et al. (1996) found a similar result for ringed seals. Validation of the growth layer deposition rate for specific species has been done using teeth from known-age animals (Hohn et al., 1989) or teeth from animals that had been exposed to tetracycline at a known point in time. Tetracycline binds with calcium and is incorporated into active tissues (e.g., teeth and bone) within 48 hours of administration (Frost, 1983). Under visible light, tetracycline-marked bone and/or teeth exhibit yellow brown coloration. Under fluorescent light, marked bone exhibits a yellow-gold fluorescence.


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Age Protocol

Teeth are the best tissues to be collected from odontocetes and pinnipeds. For small odontocetes, it is standard to collect teeth from the middle of the left mandible; six to eight teeth should be collected if the skull will not be kept. When the left mandible is not available, center teeth from the other mandible or from the maxilla are satisfactory, with the emphasis being on large, straight teeth. For pinnipeds, the best tooth to collect may depend on the relative age of the animal (juvenile, adult, old adult). To be certain that an accurate age can be obtained may require collecting several teeth, including canines and post-canines. For manatees and large whales, the ear bones should be collected. Because growth layers are integral to teeth and bone, these tissues are not sensitive to most means of storage. They can be frozen in plastic bags or vials, stored in 70% ethanol, or cleaned of soft tissue and dried. Short-term storage in formalin is acceptable. They also can be soaked in water to facilitate cleaning prior to preservation or further analyses. Care should be taken that the teeth are not damaged or broken during extraction. In certain field situations, it may be more practical to collect and save the entire mandible or skull with teeth intact for later extraction and processing. If earplugs can be collected, they should be handled gently (because they are fragile), and fixed in formalin (Lockyer, 1984). For estimation of physical maturation, physeal fusion of bones, such as vertebrae or carpal/metacarpal bones, may be evaluated from frozen or dried samples. Radiographs of flippers may assist with maturation determination, and whole flippers can be frozen for later examination. Eyes should be collected and frozen for extraction and analyses of the lenses from condition code 2 animals (George et al., 1999). Claws can be frozen or kept dry.

Reproductive Status How results are interpreted often is dependent on the reproductive status of a specimen. The primary question is whether a specimen is sexually mature or not. Then, for a mature female, the next issue is whether the animal is pregnant, lactating, or resting (not pregnant or lactating). Presence of milk in the mammary glands is diagnostic of lactation, in the absence of pathology. Presence of ovulatory scars (corpus luteum or corpus albicans) is indicative of a sexually mature female. Presence of a corpus luteum is indicative of recent ovulation and, although not diagnostic of pregnancy, signals the need to examine the uterus carefully for a possible fetus. Because pinnipeds exhibit delayed implantation, it is also important to examine the uterus for presence of a blastocyst. It is helpful to know whether a mature female has actually been pregnant. In some cases, gross examination of the uterus will show if the uterus or uterine horns have been distended at some time in the past by the presence of a large fetus. Histological sections can ascertain changes to the uterus due to pregnancies not carried to term, or for a long duration. It is also helpful to know how many times a female may have been pregnant. For cetaceans, the corpora albicantia are persistent. These structures are counted in gross examination of sections from intact ovaries as a measure of the maximum number of pregnancies possible. In all cases, both ovaries should be examined to ensure no ovulations are left uncounted. Small cetaceans tend to ovulate only from their left ovary, at least until they become older adults. Maximal follicle diameter in ovaries yields data on seasonality of ovulation and whether an immature female is close to maturation. For males, confirmation of sexual maturity requires ascertaining the presence of spermatozoa either through sperm smears from fresh epididymides or from histological examination of testis and epididymis tissue. When these data are collected in concert with data on testicular size (length, width, depth, and mass) for a large number of animals, it becomes possible to bracket the size of testes from mature and immature animals. Intermediate sizes of testes still



457

must be examined histologically to determine whether the animal was pubertal or was mature with recrudescent testes because it was not breeding season. Gonads can yield important information about whether an animal was sexually mature or not, even when they are in an advanced state of decomposition. Of course, histological studies will not be possible, but the presence of corpora in females and the large change in size between immature and mature testes, testis mass, and length in males should be sufficient in most cases to classify the specimen as mature or not when examined by an experienced life-history biologist. A good rule of thumb is that if the gonads can be found, they should be collected. Reproductive Status Protocol

Gonads should always be collected even when decomposition is advanced. When possible, fresh weights and measurements should be taken. For ovaries, a measurement of mass is sufficient. Both ovaries need to be collected, especially for small cetaceans that have unsymmetrical ovulatory patterns. Whole ovaries should be fixed in 10% buffered neutral formalin when possible or frozen if no fixative is available. They should not be cut or subsampled for histology until after they have been examined in gross (whole and thick sections) for corpora. Gross examination of the uterus is performed for detection of pregnancy and whether the uterus appears to have been distended sufficiently to suggest that a pregnancy has occurred in the past. In small animals, the uterus with ovaries can easily be preserved intact in 10% buffered neutral formalin, but care should be taken that the uterus is fixed in a natural position rather than folded into a small container. If the uterus is large, it should be weighed (when possible), measured, and examined. Gross examination and measurements should include myometrial wall thickness, cervix, internal diameter of uterine horns, length of uterine horns, any lesions, fetal presence-size position, parasites, and associated lymph nodes. Representative tissue samples should be collected in 10% buffered neutral formalin. Testes and epididymides should be removed intact. If possible, testes and epididymides should be weighed separately. Testis length (not including the epididymis), width, and depth are important parameters, with mass and length of primary importance especially if the testis cannot be collected whole. Because studies have shown no significant difference in size between the left and right testes, it is not necessary to collect both testes. The opposite testis can be used to collect a subsample for histological examination, fixing all tissues in 10% buffered neutral formalin. When whole testes cannot be collected, an alternative is to collect a complete 1 cm thick cross section from one testis and a complete longitudinal section from the other, and fix these sections in formalin, preferably in a flattened position. They can later be rolled for storage in a jar. Samples from any reproductive tract lesions are also fixed in 10% neutral buffered formalin. From fresh animals, serum, feces, and urine may also be obtained. These can be used to determine reproductive hormone levels to correlate with actual physical findings. Urine, feces, or serum should be frozen for shipping and evaluation.

Pathology—Gross Necropsy Examination Gross examination of carcasses can provide valuable information for further analyses (Bonde et al., 1983; Geraci and Lounsbury, 1993; Dierauf, 1994). Photographs and written descriptions of what is seen on gross examination may often be the only documentation on some stranded animals. Gross descriptions are extremely valuable for histological interpretation, and these descriptions are submitted to the pathologist along with level A data (basic minimum data, including investigator’s name and address, source of sample, species, date, girth, weight, condition) from stranded animals (Geraci and Lounsbury, 1993) (see Chapter 4, Stranding Networks).


458


Human Interactions Although forensic techniques have not been used for examinations of marine mammal mortalities as much as they have for terrestrial animal mortalities, the use of the same principles will increase the strength of any evidence of anthropogenic cause, and improve the chances of successful management or enforcement actions. All sample collection and examination efforts, whether from a research perspective or an enforcement perspective, will benefit from careful documentation and strict protocols. The determination of cause of death in most marine mammal strandings is difficult, because autolysis often obscures much of the evidence. Developing physical criteria for the determination of some anthropogenic causes of death is, therefore, important (Kuiken, 1996; Read and Murray, 2000). Three human activities are frequently found as causes of mortality in marine mammals: gunshot, fishery interaction, and ship/boat strike. Several recent publications have described the evidence of these activities that can be determined from marine mammal carcasses (Wells and Scott, 1997; Read and Murray, 2000). The evidence obtained in the field through careful photographs, examinations, and specimen collection may be used to determine initial cause of death; however, further analyses or interpretations will be needed to finalize actual findings. Careful documentation and descriptions are required throughout the investigation and examination. In gunshot cases, the carcass can be radiographed to determine the number and position of the projectiles, and then carefully dissected to determine post-mortem vs. ante-mortem shooting, and to trace the tracks of the bullets. Bullets that are retrieved from gunshot carcasses should be washed under cold water to remove blood and tissue, carefully dried, wrapped in soft tissue paper, and packaged in a small, crushproof, labeled container (Adrian, 1996). Photographs of the necropsy findings should be permanently identified and kept with the necropsy report. In the cases of fishery interactions, photographs, measurements, and careful gross and histological examinations will provide evidence if gear is not present. Performing the forensic examination in a standardized manner will enhance the detection of fishery interactions in fresh carcasses. Care should be taken to differentiate post-mortem scavenger and autolytic changes from lesions caused by fishery entanglements (Read and Murray, 2000). Boat strikes often have pronounced gross lesions, such as propeller cuts; however, evidence can be subtle. Manatees and large whales often have fractured bones from ship strikes, but these may not be evident in superficial examination (McLellan, pers. comm.). If ship strike is suspected in large whales, the whole carcass, including bones, should be examined (Blaylock et al., 1995). Finally, all tissues and data collected in cases that may involve direct human interaction are managed through documentation of chain of custody (see Figure 1) and all evidence is stored (e.g., locked safe) so that it cannot be tampered with.

Histopathology The histological examination of tissues, including biopsies, can provide insights into normal microanatomy, morphological changes associated with disease, and evidence of anthropogenic impacts. These findings can lead to etiological diagnoses and/or causes of death. Histopathology is a critical part of the overall assessment of any marine mammal and an integral part of the assessment of the health of wild populations. To maximize the information gained, tissues should be collected as soon as possible after death. There should be a standard set of tissues that are collected and examined as part of a standard evaluation (Figure 2, Histopathology Checklist). In addition, tissues should be collected from lesions to determine cause, and from areas of injury related to human activities, such as ship strikes or gunshot wounds, to determine if the injury occurred before or after death. Histopathology can be important in assessing time


459


Lung Trachea Heart Aorta Pulmonary Artery Thymus Salivary Gland Thyroid Tonsil Tongue Esophagus Stomach Duodenum Jejunum Ileum Colon Pancreas Spleen Liver Gallbladder Adrenal Kidney Ureter Urinary Bladder Urethra

Gonad Prostate Uterus Penis Eye (L/R) Brain Spinal Cord Bone Marrow Muscle Skin Blubber

Lymph Nodes: Submandibular Cranial cervical Prescapular Axillary Tracheobronchial Hilar Gastric Hepatic Mesenteric Colonic Sublumbar Inguinal

OTHER: Figure 2 Histopathology Checklist. In performing gross necropsy examinations, an attempt should be made to obtain each of the tissues on this list. Using the check boxes will help both the investigator and the pathologist who receives the samples.

of injury relative to death in animals that have fractures or wounds possibly due to ship or boat strikes. Even in cases of animals with obvious evidence of human interaction, important information on disease, morphology, and basic biology can be obtained through standard histopathological examination. Standardized necropsy and specimen collection forms should be used by the examiner (Geraci and Lounsbury, 1993; Dierauf, 1994), as good necropsy descriptions and history will assist the pathologist in histological interpretation and diagnosis. Pathologists must have experience with examination of tissues from marine mammals. Histopathology Protocol

Standard protocols for collection of tissues for histopathological examination are used. Tissues collected should be no larger than 3 × 3 cm and ideally 0.5 cm thick. If larger samples are collected, numerous parallel cuts should be made in the tissue to improve fixative penetration (Geraci and Lounsbury, 1993). For standard evaluations, all tissues should be preserved in 10% buffered neutral formalin at a ratio of 1:10, tissue to fixative. For specific studies, other fixatives may be preferred, but maintaining standard histological fixation must become routine for marine mammal mortality investigations. All tissues from the same animal can be placed in the same container; however, specific lesions should be tagged or placed in labeled cassettes for identification. For each case, two labels should be used, one inside the container and one outside the container, and each container should only contain one case. Tissues should be allowed to fix for at least 48 hours before shipping. There are specific requirements for shipping tissues in formalin, since formalin is considered a hazardous substance.

Acoustic Pathology The assessment of damage to hearing and vestibular structures is increasing in importance (see Chapter 1, Sentinels), as management entities strive to determine the impacts of anthropogenic


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acoustic sources and activities on marine mammals (Gisner, 1998). Examination and collection of specimens from hearing structures is important, both in those cases in which acoustic trauma is suspected and in normal strandings to develop baselines. Understanding hearing parameters for various species is critical to the assessment of the potential impacts of anthropogenic noise on marine mammals. Post-mortem examinations to evaluate acoustic integrity include computerized tomography, gross dissections, and histological examinations (Ketten et al., 1993; Ketten, 1997). Acoustic Pathology Protocol

Ear extraction must be performed by skilled personnel who possess knowledge of the anatomy of marine mammals. If one is unfamiliar with these techniques, the best approach is to freeze the whole head for later extraction and examination. If the animal is small, the head may be frozen intact and shipped to a laboratory for computerized tomography, ear extraction, and dissection; however, some gross and histological parameters may be lost, making some interpretations more difficult. If frozen, then the head may be thawed in fixative rather than in water or in air, to decrease autolytic changes. When possible, the ears should be collected fresh in the field from a ventral approach according to the method of Ketten (Blaylock et al., 1995). The skull is positioned with its dorsal surface down, the mandible and associated soft tissues carefully examined and removed, and a knife or chisel used to remove the tympano-periotic complex. Care should be taken not to fracture the periotic complex. This complex should be placed in 10% buffered neutral formalin and maintained in formalin for at least 1 week prior to shipping. Fixative may be injected into the round window in fresh specimens to ensure rapid fixation. In addition to examination of the ears, the rest of the head should also be carefully examined and tissues collected for assessment of damage to acoustic fats, auditory canals, eyes (retina/sclera), and other soft tissues. Examination of other body compartments will assist with the diagnosis and interpretation of findings. Protocols for specimen collection for life-history data are listed in Table 2.

Infectious Diseases Bacteriology The collection and analyses of specimens to determine bacterial flora in marine mammals are done routinely in live capture examinations, and may be done in some carcass examinations. Establishing the background or historic flora for a species, stock, or individual animal, along with assessing any associated lesions, is important in the interpretation of mortality events, disease outbreaks, and diagnosis of disease in the individual animal. Bacteriology Protocol

Samples should be collected as aseptically as possible and as early in the necropsy as allowable. Surfaces can be sterilized by searing, or disinfected by wiping with 10% neutral buffered formalin or 70% ethanol and allowed to air-dry. Whole tissues or tissue pieces collected should be large enough to allow for trimming in the laboratory. These tissues can be frozen (−70°C) for shipment or storage. Swabs (with appropriate bacterial transport media) may be collected from external lesions, external orifices, or from fluid-filled cavities. These should be refrigerated and processed as quickly as possible after collection. Fluid samples can be collected with a syringe and needle through a cleaned surface (see above). The fluid should then be placed in anaerobic and/or aerobic transport media. With more advanced decomposition, bone marrow may be used, since bone marrow contamination due to autolysis is slower (Geraci and Lounsbury, 1993). If sterile bone biopsy tools are available, they should

Pinnipeds Mysticetes Mysticetes All Otters

Cetaceans

Both ovaries, samples from both testes, other organs as noted

Lens Claws Stomach

Premolar (post-canine) Canine, incisor Canine Canine Mandibular canine First premolar, either jaw First premolar, either jaw Tympanoperiotic bone (periotic dome) Tympanic bullae; vertebrae; metacarpals Metacarpals

Phocids Otariids Sirenians (dugongs) Odobenids Ursids Otters Sirenians (manatees)

Mandible (left)

Varies with tissue used

Collection Site

Odontocetes

All

Taxonomic Group

1–5

1–4

2–5 2 2–5 2 1–4 1–3

2–5

2–5 1 2–5 2–5 2–5 2–5 2–5 2–5

1–5

1–5

Code

Whole ovaries; whole testes with epididymis or full cross and longitudinal sections of testis; whole or portions of uterus

Whole Whole Whole volume of stomach contents when possible

Whole intact

Take morphometrics before fixing

10% buffered neutral formalin Dried Frozen Whole in 10% glycerin in 70% ethanol Piscivores—Freeze intact Herbivores—10% neutral buffered formalin 10% neutral buffered formalin in normal position; can be frozen if no fixative available

10% buffered neutral formalin, or freeze

. 70% ethanol or freeze whole

5–20% DMSO solution; 80% EtOH, saturated salt; or freeze

1 × 0.5 cm, cut in strips, whole bone or teeth; 10–20 ml of whole blood

Whole tooth with root intact; whole jaw

Fixative

Size


Morphometrics

Reproductive status

Prey

Earplugs Baleen Eyes Claws Stomach contents Feces Gonads Uterus Serum

Bone

Epidermis Muscle Leukocytes Bone Teeth Teeth

Genetics

Age

Sample

Analysis

TABLE 2 Protocols for Specimen Collection for Life-History Data


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be used; however, collection and freezing of whole small bones is also acceptable. Be sure that the bone collected has marrow. It is preferred that bacteriological samples be processed immediately, but if that is not possible, they should be frozen at −70°C.

Virology A systematic approach is necessary to detect diseases caused by viruses. The routine collection of specimens for detection of viruses may be done in fresh carcasses to determine viral baselines in marine mammals. In mortality events or disease outbreaks, such collections are even more critical for determining the cause(s) of the mortalities. Routine evaluations with established baselines and established laboratory working relationships are essential for such investigations. Virology Protocol

Cultures, polymerase chain reaction (PCR), serology, and electron microscopic evaluations can be used for assessment of viruses in marine mammals. Although the focus of each collection is often a specific target organ, there may be cases in which the site of the potential infection is not obvious. In such cases, several organs (including lymph nodes, spleen, blood, or a targeted system) should be collected. Since lymph nodes are not described for most marine mammals, lymph nodes in any areas of the lesions should be collected, along with lung- and gut-associated lymph nodes. Samples for viral culture should be collected as aseptically as possible from the target tissue(s). Tissue specimens or swabs are placed in viral transport media or in 1 to 2 ml of physiological saline with 5% bovine serum albumin containing approximately 50 µg/ml of gentamicin, and shipped immediately (Castro and Heuschele, 1992). If immediate shipment or culturing is not possible, then tissues may be frozen at −70 to −80°C or colder for later shipment and isolation. Samples collected and frozen should be large enough to allow for trimming and subculturing in the laboratory (6 3 3 cm ; ∼2 in. ) from large organs or whole small organs (adrenal, lymph nodes) (Geraci and Lounsbury, 1993). Ideally, samples for electron microscopy should be collected in 3% gluter3 aldehyde as small (1 mm ) pieces; however, samples may be taken from formalin-fixed tissue samples. PCR or nucleic acid hybridization can be performed on either frozen or fixed samples, although fresh-frozen samples are preferred (−70 to −80°C). Serum should be collected when possible for evaluation of serum antibody titers (see Chapter 15, Viral Diseases).

Parasitology Collection and examination of parasites are important from both health and life-history perspectives. The types, age class of infection, and quantity of parasites may provide insights into the feeding ecology or stock, as well as disease, condition, and health status of the animal. Documentation (written and photographic) of the location of the parasites, parasite numbers, and types of lesions associated with the parasites will assist with the interpretation of the general health condition of the animal. Voucher specimens or representative samples of each type of parasite seen during examination will assist with accurate identification and interpretation. Parasitology Protocol

Several references for the collection and preservation of parasites are available (Geraci and Lounsbury, 1993; Eros et al., 2000) (see Chapter 18, Parasitic Diseases). Ideally, parasites should be collected intact and fresh, and any associated lesions should be collected in 10% buffered neutral formalin. Protocols for specimen collection for detection of the presence of infectious diseases are listed in Table 3.

Serology

Electron microscopy (EM)

Polymerase chain reaction (PCR)

Culture

Test Tissue, blood, fluid, swabs of lesions Tissue, blood, fluid Tissue/ concentrated fluid Serum, vitreous humor

Sample

Any target tissue or lesion Venipuncture, heart blood, eye

Any lesion, target organ, or blood

Any lesion or target organ

Site to Sample

TABLE 3 Protocols for Specimen Collection for Infectious Diseases

1–2

1–2

1–2

1–2

Code 3

>5 ml

1 mm tissue sample

3

6 cm tissue sample, swab from lesion, 1 ml fluid Collect sterile samples 3 6 cm tissue sample

Sample

Frozen at −80°C after serum is separated

Fixed in 3% gluteraldehyde

Fresh frozen sample (−80°C)

Transport media or frozen (−80°C)

Storage

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Non-Infectious Diseases Toxicology Chemical pollutants are often considered to cause or contribute to mortality events and illnesses, since marine mammals have been shown to accumulate high levels of persistent organic pollutants and some elements (O’Shea et al., 1999). Samples are collected for analyses of persistent organic pollutants, which include compounds of both current and historic use. Other compounds in current use include nonpersistent compounds, polyaromatic hydrocarbons (PAHs; Krahn et al., 1993), and essential and anthropogenic elements (see Chapter 22, Toxicology). There have been numerous studies evaluating the tissue residue levels of persistent organic pollutants, but fewer studies have evaluated biomarkers of effects or lesions associated with pollutant loads (O’Shea et al., 1999) (see Chapter 22, Toxicology). To examine the potential impacts of pollutants on health, collections of tissues for biomarker analyses, as well as general pathology and disease assessment, should be performed on any animal for which contaminant samples are collected. In addition, numerous studies have shown that age and sex significantly affect tissue residue levels for most compounds; therefore, life-history sample collections and analyses are essential for accurate interpretation of tissue residue data. Toxicology Protocol

For assessment of tissue levels of persistent and nonpersistent organic pollutants that are lipophilic, blubber, milk, blood, and liver are the tissues typically analyzed; for assessment of elements, kidney, liver, blood, and skin (epidermis) may also be analyzed. However, target organs, if known, should also be collected for complete evaluation of impacts and residue levels in marine mammals. Blubber is collected from specific sites (depending on the taxon) and samples should be full thickness, since some species have both vertical and horizontal stratification in blubber. For real-time monitoring or assessment, a minimum of 20 g (ideally 100 g) should be collected of each tissue type with a clean stainless-steel knife. Tissues are collected in clean glass jars or in Teflon bags and stored at temperatures less than −80°C. For a limited time, tissues can be stored at temperatures greater than −80°C; however, if long-term storage is expected, the tissues should be stored at −80°C and, if tissues are to be archived, the tissues should be stored in liquid nitrogen. Degradation of the tissues continues at storage temperatures warmer than −80°C. When collecting tissues, ensure that the specimens or collecting instruments are not in contact with aerosols of insect repellent, smoke, exhaust fumes, petroleum fumes, or other chemical contaminants that may alter the chemical analyses of the tissues. Tissues may also be contaminated during the necropsy by gut contents or blood, thereby altering the actual measured values. Whole blood, serum, or plasma has been used for chemical analyses; a minimum of 10 ml of selected matrix should be collected and stored frozen in clean glass jars or Teflon jars/bags. Because storage of tissues or fluids in plastic can alter the chemical analyses for some compounds, tags and collection forms must note the use of such plastics. Whenever tissues are collected for pollutant analyses, a field collection description should include the conditions under which the tissues were collected and the materials used for collection, processing, and storage. Often investigators request collection of tissues for the assessment of PAH exposure or effects. Since PAHs are rapidly metabolized in marine mammals, the likelihood of finding circulating levels or tissue levels indicative of acute exposure are low. However, both serum and liver can be assessed in acute exposure cases. Some researchers use the collection of tissues (dermis or liver) for assessment of cytochrome P-4501A as a surrogate for PAH exposure; however, several



465

compounds increase levels of cytochrome P-4501A (see Chapter 22, Toxicology). An elevation in enzyme does not always equate to an elevation in exposure to PAH. Liver or skin should be collected and fixed or frozen for analysis of cytochrome P-4501A. In marine mammals, the tissue of choice for collection to detect exposure to PAHs is bile (Krahn et al., 1993). From those taxa that have gallbladders, collection of bile is performed by withdrawing fluid from the gallbladder using a syringe or by clamping off and excising the gallbladder. Once the gallbladder is excised, bile may be poured into a dark, cleaned glass container. This is the preferred collection method for pinnipeds. Collection of bile from cetaceans is more difficult, but can be accomplished by withdrawing bile from the large hepatic duct. Bile should be protected from light and can be placed in dark jars or in clear glass containers that have been wrapped in foil. These containers should be stored frozen at −80°C. Select tissues from some animals should be stored in a local archive, in the National Marine Mammal Tissue Bank (NMMTB), or in the Alaska Marine Mammal Tissue Archival Project. When collecting tissues for archiving, strict protocols must be adhered to and detailed documentation should accompany each specimen. The NMMTB is a cooperative program that collects specimens from marine mammals and archives them at the National Biological Environmental Specimen Bank at the National Institute of Standards and Technology (NIST). Protocols and selection of animals have been published (Geraci and Lounsbury, 1993). Only tissues from select fresh (code 2) animals are included in the NMMTB. For inclusion in the NMMTB, a minimum of 400 g of liver, kidney, and blubber should be collected as cleanly as possible, trimmed utilizing a titanium knife on a Teflon surface, and placed in Teflon jars or bags. The tissues are homogenized and stored in liquid nitrogen for future retrospective studies. Tissues are available for use through application to the National Marine Fisheries Service, Office of Protected Resources. Analytical quality assurance is important for all analyses but has been of particular emphasis for analyses of chemical pollutants. The Marine Mammal Health and Stranding Response Program has established an analytical quality assurance component of the program with the NIST. The analytical quality assurance component was designed to ensure the accuracy, precision, level of detection, and intercomparability of data resulting from chemical analyses of marine mammal specimens. The program consists of interlaboratory comparison exercises and preparation and development of marine mammal tissue/blood controls and standard reference materials (Wise et al., 1993; Wise, 1993; Schantz et al., 1995). Other international programs have interlaboratory or control materials. Laboratories that perform chemical analyses in marine mammal tissues are encouraged to participate in these ongoing programs. Protocols for specimen collection for detection of chemical pollutants are listed in Table 4.

Harmful Algal Blooms Unicellular microalgae are critical members of marine food webs, and therefore many algal blooms can be beneficial to marine ecosystems. Of the 5000 species of extant marine phytoplankton, a few dozen species are known to produce potent biotoxins that enter the food chain. These harmful algal blooms and their biotoxins are increasingly recognized as causes for mortality or illness in marine mammals (see Chapter 22, Toxicology). Most algal toxins that are currently known to impact marine mammals are highly potent neurotoxins. In many cases, symptoms of intoxication are not unique, making diagnosis difficult. Furthermore, the high potency of algaltoxins results in clinical symptoms or pathologies at very low concentrations. This often makes confirmation of toxin presence somewhat problematic. Certain biotoxins are rapidly cleared through the urine and feces, and may not be detectable

Specimen

Blubber Liver Brain Blood Other target organs

Bile Liver Blood

Kidney Liver Skin (epidermis in cetaceans, skin in all others) Blood Target organ

Type of Analysis

Organochlorines

Polyaromatic hydrocarbons

Elements

Left caudal lobe of liver, left kidney, skin from left lateral wall, whole blood

Excise gallbladder by clamping off cystic or bile duct; pour bile into container

Pinnipeds Otters Sirenians 1–2, 3

1–2 early

1–2, 3

Code

A minimum of 20 g

50 g tissue 5 ml bile

Minimal 20 g; 100 g optimal for real time; 400 g for archival; >6 ml blood

Amount

Whirlpak bag or in Teflon bag, freeze at −40°C Collect with stainless steel-knife or scalpel

Frozen (liquid nitrogen) in cleaned container; protect from light; collection should be performed as soon as possible after death; deterioration is rapid

Frozen in clean glass jars or Teflon jars/bags Minimize contamination of the sample after collection

Storage

466

All

Collect bile from hepatic duct with syringe

Blubber or cutaneous fat: lateral thorax—full thickness Liver—left caudal lobe

Collection Site

Cetaceans

All

Species

TABLE 4 Protocols for Specimen Collection for Chemical Pollutants





467

in tissues even though clinical signs may be evident. For example, domoic acid is cleared from serum in as little as 2 hours in experimentally treated monkeys and rats (Truelove and Iverson, 1994). Thus the analysis of serum from California sea lions (Zalophus californianus) impacted in a 1998 domoic acid–related mortality event did not always show the presence of domoic acid, whereas urine and feces were more informative even though the animals showed classical domoic acid poisoning symptoms or death (see Chapter 2, Emerging Diseases; Chapter 22, Toxicology). Harmful Algal Bloom Protocol

For the reasons noted above, in addition to the collection of samples from affected animals, it is also useful to collect samples of prey species and of water for identification for the presence of harmful algal species (Table 5). For algal species identification, plankton samples are collected with a plankton net in a vertical tow and samples preserved in Lugol’s solution or in 2% gluteraldehyde. (To prepare Lugoll’s solution dissolve 10 g potassium iodide in 100 ml distilled water; add 5 g crystalline iodine, then 10 ml glacial acetic acid; add enough Lugoll’s Solution to water sample to make “tea-colored.”) If a plankton net is not available, whole-water samples (1 l) may be collected, preserved as noted above, and stored refrigerated (not frozen). For toxin analyses in whole water, surface water should be collected in two 1-l bottles, protected from light, and stored refrigerated. The following tissues should be collected from impacted marine mammals or large marine mammal prey species: serum, urine, stomach contents, feces, liver, kidney, brain, and lung. All tissues and fluids can be stored frozen until analyzed. Separation of serum from blood and minimal hemolysis are crucial for some detection methods, as hemoglobin can cause false-positive results. Urine and feces have proved to be the most informative fluids for a number of toxin classes and, therefore, should take priority when available. A number of rapid assay methods are available for all algal toxin classes. Generally, a rapid assay is desirable to provide a quick answer, followed by a more rigorous analytical method such as high-performance liquid chromatography–mass spectroscopy (HPLC-MS) for chemical confirmation. In addition to analytical biotoxin detection, immunoperoxidase or immunocytochemical methods are available for certain toxin classes (brevetoxin, domoic acid), which can be performed on fixed or frozen tissues (Bossart et al., 1998). Tissues should be collected from target organs or lesions. Target organs for immunocytochemical techniques are brain for domoic acid and brevetoxin and respiratory tract lesions and lung for brevetoxin. Tissues should be collected fresh using normal histological protocols (see p. 459).

Conclusions The science of marine mammal forensic medicine and knowledge of marine mammal disease, analytical methods, specimen collection, research, and management needs are growing and changing rapidly. In addition, the circumstances of various marine mammal strandings or mortality events can dictate the actual protocols used for examination. The authors recommend that for stranded marine mammals, the examiner regularly check with the national, regional, local, and/or species stranding coordinators for updated specific protocols.

Acknowledgments The authors thank Sentiel Rommel and Rebecca Duerr for comments on this chapter, and stranding network members for working with and developing protocols for improving the understanding of marine mammals.

Ciguatoxins Gambiertoxins

Okadaic acid Donphysistoxin

Donophysis spp.; Prorocentrum lima; Prorocentrum concavum

Reef fish (gonads, viscera, liver, flesh) Clams Mussels

Fish Shellfish Aerosols Water

Clams Mussels Zooplankton Fish Water Mussels Clams Fish Water

Vector

Liver Kidney

Liver Kidney

Respiratory tract Liver Blubber Serum

Kidney Urine Serum Feces

Stomach contents Liver

Tissue/Fluid

RIA MBA HPLC CT CT HPLC ELISA MBA

RBA HPLC IP

MBA RBA ELISA RIA HPLC RBA HPLC MS IP

Analytical Procedure

Fat

Fat and water

Fat

Water

Water

Solubility of Toxin

Minimum 50 g of tissue or contents into plastic bag or bottle

Minimum 50 g of tissue or contents into plastic bag or bottle; 5–10 ml of serum, whole blood, or urine; brain sections fixed for IP Minimum 50 g of tissue or contents into plastic bag or bottle; 5–10 ml of serum; respiratory or mucosal sections fixed for IP Minimum 50 g of tissue or contents into plastic bag or bottle

Minimum 50 g of tissue or contents into plastic bag or bottle

Collection

Key: CT = cellular toxicity; ELISA = enzyme-linked immunosorbent assay; HPLC = high-performance liquid chromatography; IP = immunoperoxidase; MBA = mouse bioassay; MS = mass spectroscopy; RBA = receptor-binding assay; RIA = radioimmunoassay.

Ciguatera fish poisoning (CFP) Diarrhetic shellfish poisoning (DSP)

Brevetoxins

Domoic acid Isodomoic acid Domoilactones

Saxitoxin Neosaxitoxin Gonyaitoxin Decarbamoyltoxin

Toxin

468

Gambierdiscus toxicus

Nitzschia spp.; Pseudonitzschia australis; Pseudonitzschia spp. Gymnodinium breve

Amnesic shellfish poisoning (ASP)

Neurological shellfish poisoning (NSP)

Alexandrium spp.

Organism

Paralytic shellfish poisoning (PSP)

Disease

TABLE 5 Protocols for Specimen Collection for Biotoxins





469

References Adrian, W.J., 1996, Wildlife Forensic Field Manual, Association of Midwest Fish and Game Law Enforcement Officers, Colorado Division of Wildlife, Fort Collins, 211 pp. Bada, J.L., Brown, S., and Masters, P.M., 1980, Age determination of marine mammals based on aspartic acid racemization in the teeth and lens nucleus, in Age Determination of Toothed Whales and Sirenians, Perrin, W.F., and Myrick, A.C., Jr. (Eds.), International Whaling Commission, Cambridge, U.K., 113–118. Bernt, K.E., Hammill, M.O., and Kovacs, K.M., 1996, Age estimation in grey seals (Halichoerus grypus) using incisors, Mar. Mammal Sci., 12: 476–482. Blaylock, R.A., Mase, B.G., and Driscoll, C.P., 1995, Final Report on the Workshop to Coordinate Large Whale Stranding Response in the Southeast US, SEFSC Contribution, MIA-96/97-43, 32 pp. Bonde, R.K., O’Shea, T.J., and Beck, C.A., 1983, Manual of procedures for the salvage and necropsy of carcasses of the West Indian manatee (Trichechus manatus), PB 83-255272, National Technical Information Service, Springfield, VA, 175 pp. Bossart, G.D., Baden, D.G., Ewing, R.Y., Roberts, B., and Wright, S.D., 1998, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizootic: Gross, histologic, and immunohistochemical features, Toxicol. Pathol., 26: 276–282. Castro, A.E., and Heuschele, W.P., 1992, Veterinary Diagnostic Virology, Mosby Yearbook, C.V. Mosby, St. Louis, MO, 1–5. Dierauf, L.A., 1994, Pinniped forensic, necropsy and tissue collection guide, NOAA Technical Memorandum, NMFS-OPR-94-3, 80 pp. Duignan, P.J., House, C., Odell, D.K., Wells, R.S., Hansen, L.J., Walsh, M.T., St. Aubin, D.J., Rima, B.K., and Geraci, J.R., 1996, Morbillivirus infection in bottlenose dolphins: Evidence for recurrent epizootics in the western Atlantic and Gulf of Mexico, Mar. Mammal Sci., 12: 499–515. Eros, C., Marsh, H., Bonde, R., O’Shea, T., Beck, C., Recchia, C., and Dobbs, K., 2000, Procedures for the salvage and necropsy of the dugong (Dugong dugon), Great Barrier Reef Marine Park Authority, Research Publication, 64: 1–74. Frost, H.M., 1983, Bone histomorphometry, choice of marking agent and labeling schedule, in Bone Histomorphometry: Techniques and Interpretation, Recker, R.R. (Ed.), CRC Press, Boca Raton, FL, 37–52. George, J.C., Bada, J., Zeh, J., Scott, L, Brown, S.E., O’Hara, T., and Suydam, R., 1999, Age and growth estimates of bowhead whales (Balaena mysticetus) via aspartic acid racemization, Can. J. Zool., 77: 571–580. Geraci, J.R., and Lounsbury, V.J., 1993, Specimen and data collection, in Marine Mammals Ashore: A Field Guide for Strandings, Geraci, J.R., and Lounsbury, V.J. (Eds.), Texas A&M Sea Grant Program, Galveston, 175–228. Geraci, J.R., and Lounsbury,V.J., 1997, The Florida manatee: contingency plan for health-related events, Florida Department of Environmental Protection, Florida Marine Research Institute, Contract MR199, 101 pp. Gisner, R.C., 1998, Proceedings of the workshop on the effects of anthropogenic noise in the marine environment, Office of Naval Research, San Diego, CA, 141 pp. Hohn, A.A., and Fernandez, S., 1999, Biases in dolphin age structure due to age estimation technique, Mar. Mammal Sci., 15: 1124–1132. Hohn, A.A., Scott, M.D., Wells, R.S., Sweeney, J.C., and Irvine, A.B., 1989, Growth layers in teeth from known age free ranging bottlenose dolphins, Mar. Mammal Sci., 5: 315–342. Kato, H., 1984, Readability of Antarctic minke whale earplugs, Rep. Int. Whaling Comm., 33: 393–399. Ketten, D.R., 1997, Structure and function in whale ears, Bioacoustics, 8: 103–137. Ketten, D.R., Lien, J., and Todd, S., 1993, Blast injury in humpback whale ears: Evidence and implications, J. Acoust. Soc. Am., 94: 1849–1850.


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Klevezal, G.A., 1996, Recording Structures of Mammals: Determination of Age and Reconstruction of Life History, A.A. Balkema, Rotterdam, the Netherlands, 286 pp. Krahn, M.M., Ylitalo, G.M., Buzitis, J., Chan, S.-L., and Varanasi, U., 1993, Rapid high-performance liquid chromatographic methods that screen for aromatic compounds in environmental samples, J. Chromatogr., 642: 15–32. Krahn, M.M., Ylitalo, G.M., Burrows, D.G., Calambokidis, J., Moore, S.E., Gosho, M., Gearin, P., Plesha, P.D., Brownell, R.L., Jr., Blokhin, S.A., Tilbury, K.L., Rowles, T., and Stein, J.E., in press, Environmental assessment of eastern North Pacific gray whales (Eschrichtius robustus): Lipid and organochlorine contaminant profiles, J. Cetacean Res. Manage. Kuiken, T. (Ed.), 1996, Diagnosis of by-catch in cetaceans, in Proceedings of the Second European Cetacean Society Workshop on Cetacean Pathology, Montpellier, France, 2 March 1994, 43 pp. Lockyer, C.H., 1984, Age determination by means of the earplug in baleen whales, Rep. Int. Whaling Comm., 34: 692–696. Marmontel, M., O’Shea, T.J., Kochman, H., and Humphrey, S.R., 1996, Age determination in manatees using growth layer group counts in bone, Mar. Mammal Sci., 12: 54–58. Oosthuizen, W.H., 1997, Evaluation of an effective method to estimate age of Cape fur seals using ground tooth sections, Mar. Mammal Sci., 13: 683–693. O’Shea, T.J., Reeves, R.R., and Long, A.K., 1999, Marine mammals and persistent organic contaminants, in Proceedings of the Marine Mammal Commission Workshop, Keystone, CO, 150 pp. Perrin, W.F., and Myrick., A.C., Jr., 1980, Age determination of toothed whales and sirenians: Growth of odontocetes and sirenians; problems in age determination, in Proceedings of the International Conference on Determining Age of Odontocete Cetaceans (and Sirenians), International Whaling Commission, Cambridge, U.K., Special Issue 3, 229 pp. Read, A.J., and Murray, K.T., 2000, Gross evidence of human-induced mortality in small cetaceans, U.S. Department of Commerce, NOAA Technical Memo., NMFS-OPR-15, 21 pp. Schantz, M.M., Koster, B.J., Oakley, L.M., Schiller, S.B., and Wise, S.A., 1995, Certification of polychlorinated biphenyl congeners and chlorinated pesticides in a whale blubber standard reference material, Anal. Chem., 67: 901–910. Scheffer, V.B., 1950, Growth layers on the teeth of Pinnipedia as an indicator of age, Science, 112: 309–311. Stevick, P.T., 1999, Age-length relationships in humpback whales: a comparison of strandings in the western North Atlantic with commercial catches, Mar. Mammal Sci., 15: 725–737. Stewart, R.E.A., Stewart, B.E., Stirling, I., and Street, E., 1996, Count of growth layer groups in cementum and dentine of ringed seals, Mar. Mammal Sci., 12: 383–401. Thomson, R.B., Butterworth, D.S., and Kato, H., 1999, Has the age at transition of Southern Hemisphere minke whales declined over recent decades? Mar. Mammal Sci., 15: 661–682. Truelove, J., and Iverson, F., 1994, Serum domoic acid clearance and clinical observations in the cynomolgus monkey and Sprague-Dawley rat following a single i.v. dose, Bull. Environ. Contam. and Toxicol., 52: 479–486. Wells, R.S., and Scott, M.D., 1997, Seasonal incidence of boat strikes on bottlenose dolphins near Sarasota, Florida, Mar. Mammal Sci., 13: 475–480. Wilkinson, D.M., 1996, National contingency plan for response to unusual marine mammal mortality events, NOAA Technical Memorandum, NMFS-OPR-9, 118 pp. Wise, S.A., 1993, Quality assurance of contaminant measurements in marine mammal tissues, in Coast Zone 93: Proceedings of the 8th Symposium on the Coastal and Ocean Management, Magoon, O.T., Wilson, W.S., Converse, H., and Tobin, L.T. (Eds.), New York, 3: 2531–2541. Wise, S.A., Schantz, M.M., Koster, B.J., Demiralp, R., Mackey, E.A., Greenberg, R.R., Burow, M., Ostapczuk, P., and Lillestolen, T.I., 1993, Development of frozen whale blubber and liver reference materials for the measurement of organic and inorganic contaminants, Fresenius J. Anal. Chem., 345: 270–277.


22

Toxicology Todd M. O’Hara and Thomas J. O’Shea

Introduction A poison is any substance that, when ingested, inhaled, absorbed, applied to, injected into, or developed within the body, in relatively small amounts, may cause damage to body structure or disturbance of function (Fowler, 1993). Clinical and diagnostic toxicology utilizes a variety of techniques to determine the role a “poison” has in producing an adverse effect on health (i.e., mortality event, disease, low recruitment). This expertise is commonly used in human and domestic animal disease investigations, and is very dependent on data generated from related research activities. With respect to marine mammals, specific diagnostic toxicology expertise and supporting research are severely lacking. An understanding of chemical absorption, distribution, metabolism, and excretion is limited for most species of marine mammals. However, a pharmacokinetic model was recently proposed for hydrophobic agents, based on polychlorinated biphenyls (PCBs) and belugas (Delphinapterus leucas) (Hickie et al., 1999). Reviews of chemicals and their effects on marine mammals include O’Shea (1999), Reijnders et al. (1999), and Vos et al. (in press). However, other than chemical residue data, limited information is available, especially on effects. Marine mammals fill many diverse ecological roles, from primary consumers to top carnivores, and stem from several distinct evolutionary lines. As such, they are exposed to a wide range of types and amounts of potentially toxic substances, and should be expected to show different effects from these exposures. Thus, it may not be appropriate to generalize about toxic substances and their possible impacts across marine mammals as a group. Marine mammals also exhibit a wide range of functional and morphological adaptations that may influence susceptibility or resistance to toxic substances in ways that science has not yet unraveled. These include the largest body sizes ever evolved in animals, unusually low mass-specific metabolic rates (sirenians), physiological and biochemical adaptations for deep diving, large storage compartments (blood, lipid), and wide amplitudes of seasonal cycles in fat storage and mobilization. Arctic mammals are larger, longer lived, and relatively more lipid rich, all of which affect lipophilic contaminant accumulation (Alexander, 1995). This chapter has a chemical class–based organization, followed by a target organ or systematic review (Table 1) to help with unexplained lesions or observed affected systems. This approach relates to the fact that two major approaches for investigating contaminants in marine mammals have predominated in this field: the first is unexplained lesions or effects (e.g., population decline) with no obvious cause, implicating contaminants; the second is high levels of a contaminant(s) “seeking” a lesion or effect.

0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC

471

Mercury (daily, 25 mg/kg, 20–26 days)

PCB methyl sulfones

Brevetoxin (inhalation)

Harp seal (Pagophilus groenlandicus)

Manatee (Trichechus manatus latirostris)

Northern elephant seal (Mirounga angustirostris)

OCs

Claw deformations with dyskeratosis, hypotrichosis and alopecia, hyperkeratosis of hair follicles Extensive alopecia, hyperpigmentation, epidermal ulceration, massive skin necrosis, hyperkeratosis, squamous metaplasia, atrophy of sebaceous glands

Many

Integument, Sense Organs, Mouth

Lesion

Mild morbidity to death

None, loss of insulation, organ failure, death

Renal failure, uremia, toxic hepatitis

Liver and Kidneys

Accumulate in lung and produce cellular damage

Severe congestion, edema and hemorrhage of nasopharynx and lung (catarrhal rhinitis, multiorgan hemosiderosis)

Effect

Death

Muscle fasciculations, incoordination, loss of righting reflex (acute), followed by hemopathy (anemia, hemosiderosis) Exacerbate pathology associated with morbillivirus

Ronald et al., 1977

Troisi et al., 1997


Beckmen et al., 1997

Bergman and Olsson, 1985

See text

Reference

472

Respiratory and Cardiovascular Systems

Gray seal (Halichoerus grypus)

Organochlorines (OCs)

a

Many

Species

Oil

Chemical

TABLE 1 Documented or Proposed Effects of Contaminants on Marine Mammals (arranged by organ system)



Bottlenose dolphin (Tursiops truncatus) captive

Manatee (Trichechus manatus latirostris) Mediterranean monk seal (Monachus monachus) California sea lions (3) (Zalophus californianus)

Lead (25 g of airgun pellets)

Brevetoxin

OCs

Domoic acid

Saxitoxin (in liver and brain)

Ringed (Phoca hispida) and gray seals (Halichoerus grypus)

Manatee (Trichechus manatus latirostris)

Brevetoxin

c

Bottlenose dolphin (Tursiops truncatus) captive

Lead (25 g of airgun pellets)

b

Uterine stenosis and occlusions

Reproductive System

Lethargy, motor incoordination, and paralysis Zonal vacuolation of hippocampal neuropile, severe in ventral hippocampus

Intramyelinic vacuoles of optic nerve axons, vascular congestion of meninges, fine vacuolation of superficial cortex, white matter tract vacuolation of cerebrum, cerebellum Severe congestion of brain and nonsuppurative leptomeningitis

Central Nervous System

Elevated serum AST and GGT, and bilirubin; liver (hemosiderosis, hepatocytic fatty vacuolation), and kidney (vacuolar degeneration of cortical tubule epithelium, cortical hemosiderin deposits, and acidfast intranuclear inclusion bodies) Severe congestion of the liver and kidney

Sterility

67% of a local population died in 1997 70 clinically affected; displayed seizures, head weaving, ataxia, depression, abnormal scratching, and death

Death (liver = 3.6 ppm, kidney cortex = 4.3 ppm)

Death, progressive liver damage

Toxicology

(Continued)

Helle et al. 1976a,b; Bergman and Olsson, 1985; Baker, 1989

Costas and Lopez-Rodas, 1998; Hernandez et al., 1998 Van Dolah et al., 1997; Scholin et al., 1997; 2000; Miller and Scholin, 1998; Ochoa et al., 1998; Gulland, 2000


Shlosberg et al., 1997


Shlosberg et al., 1997


473

Harbor (Phoca vitulina) and gray seals (Halichoerus grypus)

Harbor seals (Phoca vitulina)

Polar bear (Ursus maritimus) Northern fur seals (Callorhinus ursinus)

OCs

PCBs/DDT

OCs

Negative correlation to retinol, T3 and T4 Negative correlation to retinol, T3 and T4

Endocrine System

Exostosis

Endocrine-related skull asymmetry and bone lesions, hyperadrenocorticism

Compromised cohort of pups of primiparous dams?

Beckmen, 1999

Skaare, 2000

Bergman and Olsson, 1985; Zakharov and Yablokov, 1990; Bergman et al., 1992; Mortensen et al., 1992; Olsson et al., 1994 Mortensen et al., 1992

De Guise et al., 1994

Hermaphroditism Skeleton

Fujise et al., 1998

Abnormal testis

Skaare, 2000; Wiig, 1998

Subramanian et al., 1987

DeLong, 1973

Reijnders, 1986

Reference

Martineau et al., 1994

Adrenocortical dysfunction

Reduced cub survival (Svalbard, Norway)

Abortion, stillbirths, premature pups

Lower reproductive success, likely implantation failures

Effect

Implantation

Weak negative correlation between testosterone and DDE in blood and blubber

Lesion

474

PCBs

OCs

Polar bears (Ursus maritimus) Beluga (Delphinapterus leucas) Minke whale (Balaenoptera acutorostrata) Beluga (Delphinapterus leucas)

Harbor seal (Phoca vitulina) captive California sea lion (Zalophus californianus) Dall’s porpoises (Phocoenoides dalli)

Species

PCBs

DDE

OCs

e

OCs

Chemical

TABLE 1 Documented or Proposed Effects of Contaminants on Marine Mammals (arranged by organ system) (continued)



Beluga (Delphinapterus leucas) Harbor porpoise (Phocoena phocoena) Harbor seal (Phoca vitulina) captive

Striped dolphin (Stenella coeruleoalba)

PCB 138

sumPCBs (higher concentrations) OCs

PCBs (especially coplanar forms) in blubber

OCs

Manatee (Trichechus manatus latirostris) Bottlenose dolphins (Tursiops truncatus), free-ranging male

Harbor seal (Phoca vitulina) Dall’s porpoise (Phocoenoides dalli) Harbor seal (Phoca vitulina) Beluga (Delphinapterus leucas) Harbor seal (Phoca vitulina), harbor porpoise (Phocoena phocoena)

Brevetoxin

OCs

PCBs

DDE/PCBs

PCBs

Epizootic in animals with elevated PCBs

Lower serum vitamin A, white blood cells and granulocytes; lower natural killer cell activity and lymphocyte function assays (mitogen stimulation)

Correlation with reduced immune responses (in vitro), mitogen-induced proliferation responses of lymphocyte cultures Reduced proliferative responses of splenocytes

(Continued)

Kannan et al., 1993; Aguilar and Borrell, 1994a

Brouwer et al., 1989; de Swart et al., 1993; 1994; 1996; Ross et al., 1996

Cellular immunity affected more than humoral immunity

Influenced susceptibility to f morbillivirus

Jepson et al., 1999

De Guise et al., 1998

Lahvis et al., 1995

See Chapter 12, Immunology

Schumacher et al., 1993

Reijnders in O’Shea et al., 1999 Béland et al., 1992

Subramanian et al., 1987

Brouwer et al., 1989

Died from infectious disease

Immune System (spleen, lymph nodes, blood)

Thyroid colloid depletion, fibrosis

Adrenal hyperplasia

Low estradiol

Reduced testosterone

Low retinol, T3, and T4


Toxicology 475

Polar bears (Ursus maritimus) Northern fur seals (Callorhinus ursinus)

Species IgG decreased with increasing PCB Reduced response to tetanus vaccination, and lower Ig levels for pups with higher exposure via milk

Lesion Bernhoft et al., 2000 Beckmen, 1999

Compromised cohort of pups of primiparous dams(?)

Reference

Immunosuppression(?)

Effect

Brevetoxicoses has been implicated in dolphin mortality events (1946–1947; 1987–1988) (Gunter et al., 1948; Geraci et al., 1989) and previously for manatees (1963 and 1982) (Layne, 1965; O’Shea et al., 1991). b AST = Aspartate aminotransferase; GGT = γ-glutamyl transferase. c Other possible causes (i.e., morbillivirus, Osterhaus et al., 1997). d May have occurred previously in northern fur seals and sea lions (1978, 1986, 1988, and 1992) in California, and in Mexico in dolphin and sea lions. e However, these observations were confounded. Females with impaired reproduction for any reason, particularly younger animals, lack avenues for excretion of organochlorines through lactation and can be expected to have higher concentrations. f Although other hypotheses have also been advanced to explain these differences.

a

PCBs

OCs

Chemical

TABLE 1 Documented or Proposed Effects of Contaminants on Marine Mammals (arranged by organ system) (continued)


476 CRC Handbook of Marine Mammal Medicine


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Marine mammals have probably been examined for concentrations of toxic substances in tissues more than any other group of mammals. However, because of logistical, political, and financial constraints, they are among the least studied in terms of controlled experiments, making it difficult to interpret results of such surveys. In an ideal world, diagnostic toxicology should rely on consistent experimentally determined clinical and pathological findings that can be related to amounts in tissues, but little such information exists for marine mammals. One can predict pathological findings based on known effects in other mammals, but most studies of chemical residues in marine mammals are devoid of any corresponding unequivocal studies on health effects. Prediction of effects of toxins in marine mammals is also limited by single measurements of residues. It is difficult to associate a dose and time course of exposure from residues determined at a single time point. Nendza et al. (1997) indicated the importance of determining contaminant body burden in prey species to assess exposure for risk assessment properly, and that current aquatic exposure assessments are not adequate. However, the narrative sections of this chapter will provide maximum levels of concentrations of some potentially toxic substances in tissues of marine mammals, and discuss whether or not these were associated with any toxic effects in those species. This provides a yardstick against which marine mammal specialists may measure their findings. Finally, it needs to be appreciated that the presence of potentially toxic substances in tissues of marine mammals does not constitute evidence of harm. Some of these substances can probably be found in every vertebrate animal anywhere on the planet, particularly if very sophisticated chemical analyte methodology is employed. Although there are few definitive studies that have demonstrated toxic effects of environmental contaminants specifically on marine mammals, there are strong opinions that contaminants do indeed have such effects on these species. This is based largely on the growing knowledge of impacts on other species and on weighing disparate sources of sometimes isolated evidence from marine mammals. Minute traces of some compounds act as mimics of estrogens and thus can manifest toxicity through transgenerational developmental and reproductive effects (Yamamoto et al., 1996). This is generally presented as the “endocrine disruptor hypothesis” (see Chapter 10, Endocrinology). The reader should be aware that the existence of such endocrine disruption at low environmental exposures as a widespread phenomenon (not just in marine mammals) has not yet stood the test of scientific scrutiny. A major review of this concept was recently completed by the National Academy of Sciences (National Research Council, 1999). New research should attempt to test this hypothesis, as embodied in a number of recommendations from the Keystone, Colorado workshop (O’Shea et al., 1999).

Classes of Toxicants There are numerous ways to classify chemicals, including grouping by chemical structure (e.g., polyaromatic hydrocarbons, PAHs), pharmacological or toxicological mechanism of action or consequence (e.g., cholinesterase inhibitors, carcinogens), environmental persistence or half-life (e.g., persistent organic pollutants, POPs), chemical behavior (e.g., oxidants), and by clinical use (e.g., antibiotics). This can be confusing and is obviously dependent on one’s perspective. A clinically useful chemical can easily become a toxicant when used improperly. Paracelsus (1493–1541) said, “All substances are poisons; there is none which is not a poison. The right dose differentiates a poison and remedy” (Casarett and Bruce, 1980).


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Elements Elemental interactions are critical, as many toxic nonessential elements interfere with essential elements. High levels of essential elements may be toxic, and toxicity can vary with the form of the element. O’Shea and Tanabe (in press) noted that, through 1998, the published literature contained results from determinations of concentrations of as many as 50 elements from multiple organs and tissues of over 7000 individuals in at least 64 species of marine mammals. These extensive surveys have provided a large amount of data on differential occurrences of residues in various body components, and their changes with age (Martin et al., 1976; Denton et al., 1980; Wagemann et al., 1983; Norstrom et al., 1986; Honda et al., 1987; Frank et al., 1992; Malcolm et al., 1994; Noda et al., 1995). However, there has been little evidence of toxicity or impacts on populations of marine mammals due to elements, from the limited studies conducted to date (Dietz et al., 1998). The elements of greatest concern that have been studied most intensively are mercury, cadmium, lead, and, more recently, the organotins. Other elements occur in seemingly low concentrations, or have suspected roles as essential nutrients, or lack evidence of harmful effects at concentrations reported.

Mercury Mercury (Hg) is a toxic, nonessential element that can biomagnify in food chains, particularly in its methylated form, which is the most toxic (Law et al., 1996). High concentrations of mercury are found in piscivorous marine mammals, particularly in areas high in mercury of geological origin. Mercury exposure may also occur from point sources, such as mining areas near river dolphin habitat (Rosas and Lehti, 1996). Most data on mercury in marine mammals are based on total, or inorganic, mercury, because of the expense of quantifying concentrations of the methylated form. The primary focus of these studies has been the determination of mercury concentrations in liver, which are usually higher than in other tissues. Mercury levels in marine mammal liver increase with age, although the proportion of methylmercury typically decreases with age (O’Shea, 1999; Siebert et al., 1999). An appreciation of the difficulty in measuring levels and specific forms of mercury in marine mammal tissues is warranted and must be considered when making comparisons between studies (Armstrong et al., 1999). High mercury concentrations now known to be common in livers of marine mammals were at first shocking, because they would indicate toxicoses in terrestrial mammals. Depending upon the proportion of methylmercury, high concentrations of mercury in tissues of marine mammals could pose health risks to people who consume them (Paludan-Müller et al., 1993; Simmonds et al., 1994). However, knowledge of consumption rates, presence of selenium, and bioavailability is needed to make this assessment and is beyond the scope of this chapter. Marine mammals have an apparent capacity to detoxify and store mercury. Recent reports verify earlier findings of extraordinarily high concentrations of total mercury in livers of marine mammals (Koeman and van den Ven, 1975; Smith and Armstrong, 1975; Roberts et al., 1976). Maximum concentrations reported include 626 ppm (dry weight) in pilot whales (Globicephalus spp.), 2 ppm (wet weight) in Risso’s dolphin (Grampus griseus) (Storelli et al., 1999), 751 ppm (wet weight) in harbor seals (Phoca vitulina) (Reijnders, 1980), 1097 ppm (wet weight) in gray seals (Halichoerus grypus) (Simmonds et al., 1993), and 5400 ppm (dry weight) in striped dolphins (Stenella coereloalba) (Leonzio et al., 1992). Over 13,000 ppm (dry weight) mercury was found in bottlenose dolphins (Tursiops truncatus) from the Mediterranean Sea, an area with naturally high background mercury levels of geological origin (Leonzio et al., 1992). The baleen whales and sirenians, which typically feed lower in the food chain, have lower mercury concentrations


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in liver in comparison with piscivorous species (O’Shea et al., 1984; Honda et al., 1986b; 1987; Dietz et al., 1990; Sanpera et al., 1993; Bratton et al., 1997; Woshner, 2000). Few studies have associated hepatic mercury concentrations with pathology. Rawson et al. (1993; 1995) noted mercury-associated lipofuscin-like pigment granules and liver lesions (fat globules, central necrosis, lymphocytic infiltrates) in bottlenose dolphins with relatively high concentrations of total mercury in liver (up to 443 ppm wet weight). Pigment granules were found in all dolphins with 61 ppm or more mercury; however, these animals were also the older animals in the survey. Siebert et al. (1999) determined mercury concentrations and gross and histopathological lesions in 57 harbor porpoises (Phocoena phocoena) and three white-beaked dolphins (Lagenorhyncus albirostris) from the Baltic and North Seas. No lesions indicative of mercury intoxication were found, despite liver concentrations of total mercury of up to 449 ppm dry weight, and of methylmercury of up to 26 ppm dry weight. Woshner (2000) conducted a survey for mercury-specific lesions in polar bear (Ursus maritimus), ringed seal (Phoca hispida), bowhead whale (Balaena mysticetus), and beluga tissues, and found no association of pigment levels or presence with mercury levels or histologically determined mercury distribution. The proposed tolerance of cetaceans and pinnipeds for mercury may be based on the evolution of biochemical mechanisms involving selenium. Mercury concentrations in livers that would be considered toxic in other mammals are accompanied by increased selenium levels in most marine mammals (Koeman, 1973; Koeman and van den Ven, 1975). At low tissue levels of mercury, the mercury-to-selenium molar ratio is positively correlated with the mercury tissue concentrations, but it stabilizes around a 11 molar ratio of mercury to selenium in liver at mercury concentrations of about 100 ppm (Krone et al., 1999). This 11 ratio is not, however, always evident (Woshner, 2000), and is very different from those seen in fish (Koeman, 1973; Koeman and van den Ven, 1975; Nigro and Leonzio, 1996). Mercury in fish muscle is mostly in the highly toxic methylated form (kidney and liver have higher proportions of divalent mercury), but in marine mammals the proportion of methylated mercury in livers is low, but high in muscle and epidermis (at least in cetaceans). The biochemistry of demethylation and the likely protective effect of selenium are not completely understood. They appear to involve distribution of mercury from the kidney and other sensitive organs to muscle and other tissue, competition for binding sites, formation of a mercury–selenium complex, conversion of toxic forms of mercury (methylated) to less toxic forms (i.e., divalent), and prevention of oxidative damage (Cuvin-Aralar and Furness, 1991). With low selenium levels, mercury is detoxified by binding to metallothionein proteins, a process that may cause mercury retention. Selenium apparently diverts binding of mercury away from metallothionein to higher molecular weight proteins. Deposits of mercury–selenide complexes (tiemannite) also occur in marine mammals (Martoja and Viale, 1977). Mercury–selenium correlations have been determined in tissues of several species of marine mammals (see summary in O’Shea, 1999), and results are consistent with a role for selenium in protection against mercury toxicity (Koeman, 1973; Koeman and van den Ven, 1975; Cuvin-Aralar and Furness, 1991). The gross distribution of mercury within the liver of harbor porpoises is homogenous (Stern et al., 1992), but in belugas is zonary, based on age and levels of mercury (Woshner, 2000). Within the cell, mercury and selenium occur as dense, intracellular granules located in macrophages, mainly within the liver, but also in the spleen, bone marrow, and lungs. Macrophages may accumulate mercury through phagocytosis of erythrocytes, clearing methylmercury from the bloodstream (Nigro and Leonzio, 1996). There have been a few experimental and in vitro studies of mercury toxicity in marine mammals. Some of these verified that demethylation occurs and that selenium is involved in preventing mercury toxicoses. Freeman et al. (1975) dosed harp seals (Pagophilus groenlandicus)


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with 0.25 mg/kg methylmercuric chloride daily for 2 months, and found substantial (70%) demethylation. Methylmercuric chloride was also administered to gray seals, which showed increased mercury and selenium in livers, but only increased mercury (not selenium) levels in other tissues (van de Ven et al., 1979). A two-staged excretion rate (55% with a half-life of 3 weeks, 45% with a half-life of 500 days) was calculated for a ringed seal dosed with radioactively labeled methylmercury (Tillander et al., 1972). Renal failure, uremia, toxic hepatitis, and death occurred in harp seals exposed daily to 25 mg/kg methylmercuric chloride for 20 to 26 days, but no obvious lesions occurred at 0.25 mg/kg (Holden, 1978). Harp seals dosed with methylmercuric chloride also showed a nonspecific, low level of structural damage to sensory cells of the organ of Corti, including missing or damaged stereocilia, reticular scars, and collapsed sensory cells (Ramprashad and Ronald, 1977). The implications of this damage are unknown. Alteration of gonadal and adrenal steroid synthesis was reported in harp and gray seals administered 0.25 mg/kg methylmercury (Freeman et al., 1975; Freeman and Sangalang, 1977). Splenocytes and lymphocytes of belugas cultured with mitogens were exposed to mercuric chloride solutions. Decreases in thymocyte viability and proliferative responses were observed −5 at the highest concentrations (10 M HgCl2), a level comparable with concentrations in livers of severely contaminated belugas from the St. Lawrence River (De Guise et al., 1996). However, hepatic levels of mercury are likely not achieved in the thymus. The methylated form of mercury was more potent in inhibiting cell proliferation and inducing micronuclei in in vitro cultured beluga skin fibroblasts (Gauthier et al., 1998). Cultured lymphocytes of bottlenose dolphins have greater resistance to cytotoxic and genotoxic effects of methylmercury than cells of rats or humans (Betti and Nigro, 1996).

Cadmium Cadmium (Cd) concentrations in a variety of tissues from many marine mammal species have been determined. The highest concentrations occur in kidney, where cadmium increases with age, with lesser concentrations in liver, and lower amounts in other tissues (O’Shea, 1999). Unusually high kidney concentrations have been reported in pinnipeds, such as 500 to 600 ppm (dry weight) (Anas, 1974; Wagemann, 1989; Malcolm et al., 1994), whereas 200 to 800 ppm (dry weight) or more occurs in cetaceans (Wagemann et al., 1983; 1990; Marchovecchio et al., 1990; Meador et al., 1993; Sanpera et al., 1996; Parsons, 1999), and as much as 308 ppm (dry weight) in sirenians (Denton et al., 1980). There is little difference in cadmium concentrations in tissues between the sexes, and cadmium does not readily cross the placenta (Honda et al., 1983; 1986b; Meador et al., 1993; Law et al., 1996). Elevated cadmium concentrations in marine mammals may result from naturally high cadmium concentrations in prey species (especially squid) from geological sources (Caurant and Amard-Triquet, 1995) rather than pollution (Leonzio et al., 1992; Szefer et al., 1994). Cadmium toxicosis in mammals is ameliorated by the protective action of metallothioneins. The presence of the latter has been demonstrated in pinnipeds (Olafson and Thompson, 1974; Lee et al., 1977; Mochizuki et al., 1985; Tohyama et al., 1986) and cetaceans (Wagemann et al., 1984; Goodwin et al., 1999). Other metal-binding proteins have been isolated from tissues of sperm whales (Physeter macrocephalus) (Ridlington and Whanger, 1981). Splenocytes and lymphocytes of belugas cultured with mitogens exposed to cadmium chloride solutions did −5 not decrease in viability, but at high concentrations of cadmium chloride (10 M, comparable with those in livers of belugas from the St. Lawrence River) exhibited decreased proliferation of splenocytes and thymocytes (De Guise et al., 1996). However, the relevance of this concentration to nonrenal tissue and function must be questioned, as the authors are unaware of any demonstrated


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cadmium-induced pathology in marine mammals. Humans in northern communities who consume organs from marine mammals may be at risk from high cadmium exposure, but a better determination of consumption rates and bioavailability is needed.

Lead Lead (Pb) accumulates in bone; yet most studies of lead in marine mammals have determined concentrations solely in liver, kidney, and muscle. Liver and kidney tend to have higher lead concentrations than other soft tissues; in most studies, concentrations in soft tissues are less than 1 ppm and within the normal ranges seen in other mammals. Examples of high lead concentrations reported in soft tissues of marine mammals include 11.6 to 14.8 ppm (dry weight) in livers and kidneys of pinnipeds (Goldblatt and Anthony, 1983; Warburton and Seagers, 1993), 3.5 to 4.3 ppm (wet weight) in liver and 12.4 ppm (dry weight) in kidney of odontocetes (Law et al., 1991; André et al., 1991; Leonzio et al., 1992; Kuehl et al., 1994), 2.6 to 15.9 ppm (dry weight) in liver and kidney of baleen whales (Honda et al., 1987; Parsons, 1999), 7.1 ppm (dry weight) in kidney of a sirenian (O’Shea et al., 1984), and 1.6 ppm (wet weight) in liver of a polar bear (Norheim et al., 1992). The highest concentration of lead reported in tissues of any marine mammal is 62 ppm (wet weight) in bone of a young bottlenose dolphin stranded near a lead smelter in coastal Australia (Kemper et al., 1994). Consistent effects of age and sex across species on lead accumulation have not been well established. Lead can cross the placenta, and it may be found in milk in low quantities. Lead in striped dolphin bone occurred at lower concentrations in females than in males, and lead accumulated most rapidly in calves during the suckling period (Honda et al., 1986a). There is little clinical or experimental information on toxic effects of lead in marine mammals. Shlosberg et al. (1997) examined a captive bottlenose dolphin that inadvertently ingested 25 g of lead airgun pellets and died from lead poisoning. The dolphin had elevated serum levels of aspartate aminotransferase, γ-glutamyl transferase, and bilirubin, indicating progressive liver damage. Lesions in the liver (hemosiderosis, hepatocytic fatty vacuolation), kidney (vacuolar degeneration of cortical tubular epithelium, cortical hemosiderin deposits, and acid-fast intranuclear inclusion bodies), and nervous system (intramyelinic vacuoles of the axons of the optic nerve, vascular congestion of meninges and fine vacuolation of superficial cortex, white matter tract vacuolation of the cerebrum and cerebellum) were consistent with lead poisoning. Lead concentration (wet weight) in liver was 3.6 ppm and in kidney cortex was 4.3 ppm. De Guise et al. (1996) found no significant responses in viability or cell proliferation of splenocytes and lymphocytes (from belugas) cultured with mitogens exposed to lead chloride solutions. Caution should be used when interpreting lead levels from animals killed with lead-containing projectiles.

Organotins Organotin compounds (used as marine antifoulants on boats and nets and for other industrial purposes) have recently been documented in tissues of cetaceans (Iwata et al., 1994; Kim et al., 1996; Tanabe et al., 1998; Yang et al., 1998), pinnipeds (Kim et al., 1996), and sea otters (Kannan et al., 1998). The butyltins concentrate in tissues with high protein-binding capacity, including liver, kidney, and brain (Kannan et al., 1996; 1997). Levels are typically highest in livers of marine mammals from nearshore waters and developed coastlines (Kannan et al., 1997; Tanabe et al., 1998). Total butyltin concentrations detected in livers are typically 1 to 10 ppm (wet weight) (Tanabe, 1999).


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Butyltin levels in liver of cetaceans appear to increase with age until sexual maturity, at which point they level out (Kim et al., 1996; Kannan et al., 1997). Yang et al. (1998) determined butyltin levels in beluga liver and blubber. They concluded that total butyltin in liver increased with age, levels were higher in liver than in blubber, levels varied widely (< 2 to 30 ng/g dry weight) with blubber depth (with highest levels in deepest layer), levels were higher in blubber with 90% lipid (hydrophilicity), and that levels were comparable with those of other marine vertebrates. Toxic effects of these compounds on marine mammals have been suggested, since they may be associated with immunosuppression (Kannan et al., 1997; 1998; Tanabe, 1999). In vitro studies show inhibition of P-450 enzyme activities by tributyltins in Steller sea lions (Eumatopias jubatus) and Dall’s porpoises (Phocoenoides dalli) (Kim et al., 1998). Steller sea lions had higher concentrations of butyltins in liver than in any other tissues sampled, and calculations indicate that 26% of the butyltin burden was eliminated through shedding (Kim et al., 1996). Pinnipeds have been shown to excrete organotins via shedding of the hair during molting (Tanabe, 1999). There is little evidence of significant maternal transfer through lactation or gestation (Tanabe, 1999).

Other Elements Arsenic has been reported in numerous species of marine mammals (see O’Shea, 1999), but at concentrations and/or in forms not considered toxic. Copper is an essential element that in most mammals typically decreases with age; in marine mammals it readily crosses the placental membranes, and concentrations in the fetus are usually higher than in the mother (Honda et al., 1987; Fujise et al., 1988; Muir et al., 1988a; Law et al., 1991). Copper has not been implicated as a potential toxin in marine mammals except in the case of Florida manatees; in that case such potential was suggested for specific feeding areas where copper was intensively applied as an aquatic herbicide (O’Shea et al., 1984). Selenium, which covaries with mercury, increases with age or size in liver of several species of marine mammals (O’Shea, 1999). High concentrations of silver occur in livers of Alaskan belugas, but the toxicological significance of these findings is unknown (Becker et al., 1995; Mackey et al., 1995). Tissues of marine mammals of the North Pacific have been analyzed for vanadium, which was found primarily in liver, hair (pinnipeds), and bone and which increased with age. To date, maximum concentrations found in liver are less than 4 ppm (Mackey et al., 1996).

Halogenated Organics Accumulation and Variability The greatest amount of information available in marine mammal toxicology is based on chemical analyses of accumulation of organohalogens in blubber. However, there are major gaps in knowledge about specific adverse effects of these compounds on marine mammals, as data on residue concentrations in tissues are usually not accompanied by diagnostic data on health. Target organs (i.e., central nervous system for organochlorine pesticides) are rarely sampled for chemical analyses or other types of examination, such as histopathology. This frustrating situation is due in part to logistic, institutional, and policy difficulties in developing such information for marine mammals on an experimental basis, and has even led to the observation by one leading wildlife toxicologist that “additional collection of residue data alone is not helpful and indeed, it can be argued that analytical chemical studies should only be undertaken in support of detailed biological studies” (Peakall, 1999). Nevertheless, residue


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accumulation studies have elucidated a number of important patterns, and determining sources of variability in the accumulation of these compounds is fundamental for interpreting the results of chemical analyses. The number of organohalogen contaminants identified in marine mammal blubber has increased with time and improved analytical capabilities, and new substances continue to be reported (Vetter et al., 1999). Organohalogens, particularly certain organochlorine insecticides and PCBs, are highly lipophilic. They typically reach the oceans through terrestrial runoff and atmospheric transport on particulate matter, and biomagnify in food webs as they are ingested and concentrated in the fatty tissues of organisms. In marine mammals, blubber is the main body compartment for lipid storage, and is commonly used to investigate organohalogen accumulation. In some species, blubber has been documented to contain 90 to 95% of the total body burden of organochlorines, because most of the mass of blubber is lipid, and over 90% of total body lipid is in the blubber (Tanabe et al., 1981). Total organochlorines in other organs are much lower, but correlate with tissue lipid content; thus, most organs have similar proportions of various organochlorines when expressed on a lipid weight basis. The brain usually has lower amounts because of its higher content of phospholipids, which have less of an affinity for most organochlorines (lindane is an exception). Body size is also a factor influencing residue concentrations, with the large marine mammals having higher total body burdens but lower organohalogen concentrations in blubber than smaller species (Aguilar et al., 1999). In one study, fin whales (Balaenoptera physalus) had as high as 23.5 g sumDDT (2,2-bis-(p-chlorophenyl)1,1,1-trichloroethane) (total metabolites of the insecticide DDT, see below) in their bodies, a 10- to 100-fold increase in total body burdens compared with some small cetaceans (Aguilar and Borrell, 1994b). The dynamics of storage of organochlorines in blubber is complex. Many marine mammals undergo major cyclic changes in the amounts of lipid stored in blubber. These changes correlate with seasonal fasting, breeding, lactation, and migration. Organohalogens pass into the bloodstream with lipid mobilization, but may also concentrate in the remaining blubber. The rates of these shifts are poorly understood (Aguilar et al., 1999), and can affect results of toxicological analyses and interpretation. For example, organohalogen concentrations in blubber can be diluted with rapid expansion of the amount of blubber during seasonally dependent lipid storage, whereas marine mammals found as stranded carcasses may have depleted lipid reserves due to disease or starvation, resulting in elevated organohalogen residue concentrations in blubber. Specific organohalogens that are more easily metabolized may be in lower concentrations in blubber during seasons when blubber is depleted (Weisbrod et al., 2000a). In addition, lipid content and composition of blubber vary by location on the body, and can be structurally stratified. In cetaceans, outer layers of blubber may contain a higher proportion of lipids than inner deposits, and inner deposits have more highly saturated fatty acids and may change more acutely with mobilization or deposition of lipid (Lockyer et al., 1984; Aguilar and Borrell, 1994c). Organochlorine concentrations in outer blubber layers of seals can be significantly higher than in the inner layer (Severinsen et al., 2000). Lipid composition of blubber can vary among species and can differ from that of other organs. These sources of variation require the use of standard field sampling protocols (Aguilar, 1985; 1987; Geraci and Lounsbury, 1993). Residue concentrations can also change with time when sampled post-mortem from stranded carcasses (Borrell and Aguilar, 1990). The awareness of this variation in lipid content and contaminant levels of blubber has added scrutiny to the use of biopsy techniques for accurately representing blubber contaminant burdens (Muir, pers. comm.). Age, sex, and reproductive status are other strong sources of variation in concentrations of organohalogens in marine mammals (Aguilar et al., 1999). For some organochlorines (e.g., DDT and metabolites), immature animals of either sex may have similar concentrations in blubber,


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but adult males often show significantly higher concentrations than adult females, as seen in bowhead whales (O’Hara et al., 1999). This is likely because females have an avenue of excretion, through the transfer of organochlorines to calves during both gestation and lactation. Transfer through the lipid-rich milk can be pronounced, and may have potential ramifications for the health of nursing young (Beckmen et al., 1999). Estimates of transfer through lactation are especially high for primiparous females, ranging from about 70 to 90% of the female’s total body burden of sumDDT and PCBs in odontocetes and pinnipeds (Cockcroft et al., 1989; Borrell et al., 1995; Lee et al., 1996), with lower proportions in baleen whales (Aguilar and Borrell, 1994b). Some organochlorines do not show consistent differences with age between males and females, particularly those that are more easily metabolized or are found only in lower amounts, e.g., hexachlorocyclohexane (HCHs) and hexachlorobenzene (HCBs) (Kleivane et al., 1995; Aono et al., 1997; Bernt et al., 1999). Marine mammal populations with different feeding habits may also differ notably in organohalogen contamination (Aguilar et al., 1999; Muir et al., 2000). Contamination also differs regionally according to coastal pollution inputs, with marine mammals from inshore locations near industrial or agricultural centers typically possessing higher concentrations than pelagic species or those in remote areas. The more volatile compounds, however, can be carried to remote Arctic locations through atmospheric transport (Muir et al., 1992a; Norstrom and Muir, 1994).

Organochlorine Pesticides and Metabolites Many of the organochlorine compounds are very persistent in the environment and have wellknown, experimentally determined adverse effects in mammals. Despite restrictions on or elimination of the use of many organochlorine pesticides in some developed countries, they continue to be produced and used in numerous areas of the world. Metabolites of DDT are the most commonly reported organochlorine insecticide residues found in marine mammals. In particular, p, p-DDE (2,2-bis-(p-chlorophenyl)-1,1-dichloroethylene) is typically the most abundant metabolite and its concentrations are usually far higher than DDT or TDE (DDD) (2,2-bis-(p-chlorophenyl)-1,1dichloroethane). Exceptions can occur in marine mammals in areas with very recent DDT contamination, or unusual food habits (Senthilkumar et al., 1999). The o, p-isomers of DDT metabolites are sometimes reported, usually at lower concentrations. A few studies have also reported additional metabolites of DDT in marine mammals, including methyl sulfone compounds (Bergman et al., 1994), and some studies employ ratios of DDE to DDT to estimate recency of input or degree of “aging” of DDT in ecosystems (Borrell and Reijnders, 1999). Extreme cases of contamination of marine mammals with sumDDT have resulted in concentrations of 500 to 2500 ppm or more in blubber, particularly in past decades (DeLong et al., 1973; O’Shea et al., 1980; Gaskin et al., 1982; 1983; Baird et al., 1989; Blomkvist et al., 1992). However, typical concentrations are much less than 100 ppm, with many samples at 10 ppm or less. This is particularly so in marine mammals with large body sizes or occupying low trophic levels, such as the baleen whales and sirenians, or in samples from species of the open oceans or high latitudes (O’Shea and Brownell, 1994; Ames and Van Vleet, 1996; Aono et al., 1997; Aguilar et al., 1999; O’Hara et al., 1999). Dieldrin is commonly reported in blubber of marine mammals throughout the world, but the less persistent parent compound (aldrin) and the related highly toxic form (endrin) are seldom found (Weisbrod et al., 2000b). Concentrations of dieldrin in marine mammal blubber are usually much lower than those of sumDDT, seldom reaching 10 to 15 ppm in the past, and 0.1 ppm in modern samples. The cyclodiene insecticide chlordane is a mixture of cis- and trans-isomers of chlordane, heptachlor, and nonachlor (Dearth and Hites, 1991). The more toxic heptachlor epoxide is


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the principal metabolite of heptachlor found in marine mammals (heptachlor is also used as an insecticide). Isomers of chlordane, oxychlordane (metabolite), and nonachlors, as well as heptachlor and heptachlor epoxide, have also been reported in marine mammals throughout the world, including at high latitudes (O’Shea et al., 1980; Muir et al., 1990; Norstrom and Muir, 1994; Salata et al., 1995; Aono et al., 1997; Strandberg et al., 1998; Vetter et al., 1999; O’Hara et al., 1999). Concentrations are generally low (usually

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